deliverable no 4.1: standard operating protocol (sop) on

This project has received funding from the European Union’s Horizon 2020 research and innovation programme under grant agreement No 731065

Project Title: AQUACOSM: Network of Leading European AQUAtic

MesoCOSM Facilities Connecting Mountains to Oceans from

the Arctic to the Mediterranean

Project number: 731065

Project Acronym: AQUACOSM

Proposal full title: Network of Leading European AQUAtic MesoCOSM Facilities

Connecting Mountains to Oceans from the Arctic to the

Mediterranean

Type: Research and innovation actions

Work program topics

addressed:

H2020-INFRAIA-2016-2017: Integrating and opening research

infrastructures of European interest

Deliverable No 4.1: Standard Operating Protocol (SOP) on

Sampling and Analysis of Phytoplankton, expanded for further

SOPs

Due date of

deliverable:

30 September 2017

Actual submission

date:

18 September 2017; resubmitted 29 May 2020

Version: V2.0

Main Authors: Robert Ptacnik (WCL), Deniz Başoğlu (METU), Meryem Beklioğlu (METU), Daphne

Buijert-De Gelder (NIOO), Sven Teurlincx (NIOO), Lisette N. de Senerpont Domis

(NIOO), Christian Preiler (WCL), Behzad Mostajir (CNRS), Thomas A. Davidson

(AU), Johan Wikner (UMU), Paraskevi Pitta (HCMR), Iordanis Magiopoulos

(HCMR), Aud Larsen (UNI), Sarah Fiorini (ENS)


Project ref. number 731065

Project title

AQUACOSM: NETWORK OF LEADING EUROPEAN AQUATIC MESOCOSM

FACILITIES CONNECTING MOUNTAINS TO OCEANS FROM THE ARCTIC TO

THE MEDITERRANEAN

Deliverable title Standard Operating Protocol (SOP) on Sampling and Analysis of

Phytoplankton

Deliverable number D4.1.1

Deliverable version V2.0

Contractual date of

delivery

30 September 2017

Actual date of delivery 18 September 2017, resubmission 29 May 2020

Document status Revision including addition of SOPs

Document version V2.0

Online access Yes

Diffusion Public

Nature of deliverable Report

Work package WP4.1

Partner responsible METU, WCL

Author(s) Robert Ptacnik (WCL), Deniz Başoğlu (METU), Meryem Beklioğlu (METU),

Daphne Buijert-De Gelder (NIOO), Sven Teurlincx (NIOO), Lisette N. de

Senerpont Domis (NIOO), Christian Preiler (WCL), Behzad Mostajir (CNRS),

Thomas A. Davidson (AU), Johan Wikner (UMU), Paraskevi Pitta (HCMR),

Iordanis Magiopoulos (HCMR), Aud Larsen (UNI), Sarah Fiorini (ENS)

Editor Katharina Makower (IGB), Jason Woodhouse (IGB)

Approved by Jens Nejstgaard (IGB)

EC Project Officer Agnés Robin

Abstract 1

Phytoplankton

This deliverable is a Standard Operating Protocol (SOP) that describes the

methods for sampling and analysis of phytoplankton from mesocosm

experiments carried out in all aquatic environments (fresh and marine

waters). It gathers best practice advice with a focus on sampling, counting


and other analyses of phytoplankton as well as Quality Assurance/Quality

Control (QA/QC) practices.

Use of this SOP will ensure consistency and compliance in collecting and

processing phytoplankton data from mesocosm experiments across the

AQUACOSM community, in Europe and beyond.

Abstract 2 Zooplankton This deliverable is a Standard Operating Protocol (SOP) that describes the methods for sampling and analysis of mesozooplankton from mesocosm experiments carried out in all aquatic environments (fresh and marine waters). It gathers best practice advice with a focus on sampling, counting and other analyses of mesozooplankton as well as Quality Assurance/Quality Control (QA/QC) practices.

Use of this SOP will ensure consistency and compliance in collecting and processing mesozooplankton data from mesocosm experiments across the AQUACOSM community, in Europe and beyond.

Abstract 3 Microbial

Plankton

This deliverable is a Standard Operating Protocol (SOP) that describes methods and gathers best practice advices for sampling, enumeration and other relevant analyses of microbial communities (from viruses to ciliates) from mesocosm experiments carried out in all aquatic environments (fresh and marine waters). Quality Assurance/Quality Control (QA/QC) practices are also treated.

Use of this SOP will ensure consistency and compliance in collecting and processing microbial stock abundance and bacterial activity data from mesocosm experiments across the AQUACOSM community, in Europe and beyond.

Abstract 4 Periphyton Periphyton are the algae, fungi, bacteria, and protozoa associated with substrates in aquatic environments. These organisms are a major component in energy flow and nutrient cycling in aquatic ecosystems. Periphyton and epiphyton grow on sediments and/or on surfaces of macrophytes. Especially on the surface of plants, periphyton can reduce the availability of light. In addition, periphyton is able to take up nutrients. These two characteristics may result in reduced growth of macrophytes. In this protocol, we show how to determine the light extinction by periphyton and the biomass of periphyton. This SOP follows the NIOO protocol for periphyton sampling: artificial substrate is used and it is assumed that you do not need to know the exact periphyton community composition. To this end, we use plastic strips as an artificial substrate on which periphyton can grow during a predefined period. The plastic strips remain in the water for a predefined period. After this predefined period, the strips are harvested. Subsequently, the biomass of the periphyton on the strip is quantified, as well as the light extinction caused by biofouling.

Abstract 5 Water

Chemistry

This deliverable contains Standard Operating Procedures (SOP) that describes the methods for sampling and storing samples for the analysis of water chemistry from mesocosm experiments carried out in all aquatic


environments (fresh and marine waters). It gathers best practice advice with a focus on sampling and pre-analytical processing and provides an overview of analytical methods used by project partners.

This SOP points out relevant considerations regarding planning and execution of sampling mesocosms. The listing of analytical methods used by partners allows to identify frequently used methods as well as differences in analytical procedures in the AQUACOSM community.

Abstract 6 High

Frequency Monitoring

This Standard Operating Procedure (SOP) describes methods for measuring temperature, dissolved oxygen concentration and saturation, conductivity, underwater light (PAR), pH, turbidity, depth, Chl-a, flow rate, light spectrum and nitrogen data with specific probes as well as with multi-parameter sensors in mesocosms (fresh and marine). It gathers best practice advice for field observation, cleaning, calibration, use and storage of probes as well as Quality Assurance/ Quality Control (QA/QC) measures for high frequency measurements and monitoring. In order to gather more information on the procedures for the computation of data and publication of final records, please refer to the Data-related QA/QC SOP.

Use of this SOP will ensure consistency and compliance in collecting and processing high frequency (some min to h) data from mesocosm experiments across the AQUACOSM community, in Europe and beyond.

Abstract 7 QA & QC This deliverable is a Standard Operating Protocol (SOP) that describes the methods for data quality assurance and quality control (QA/QC). It defines terms and sets out guidelines for workflow. It then describes practical processes for quality assurance and a range of tests for quality control, including suggestions for flagging systems and data handling.

Keywords • Phytoplankton Analysis, Sampling, Enumeration, Freshwater, Marine,

Algae Utermöhl

• Zooplankton Analysis, Mesozooplankton, Sampling, Enumeration,

Standard Operating Protocol, Freshwater, Brackish, Marine, Mesocosm

• Microbial Analysis, Sampling, Abundance, Growth, Activity, Freshwater,

Marine, Microscopy, Flow cytometry, Bacterial production, Isotopes,

Enzymes, Viruses, Bacteria, Flagellates, Ciliates

• Periphyton sampling, biomass quantification, artificial substrate

• Water Chemistry, Sample Storage

• High frequency measurement

• Quality assurance, Quality control, flagging


Table of Contents

I. Executive summary .................................................................................................................................. 10

II. Cross References ..................................................................................................................................... 10

III. Dissemination activities related to the Deliverable ............................................................................ 10

1. Phytoplankton ......................................................................................................................................... 11

1.1 Definitions and Terms...................................................................................................................... 11

1.2 Materials and Reagents ................................................................................................................... 12

1.3 Health and Safety Indications .......................................................................................................... 13

1.3.1 General Information ................................................................................................................ 13

1.3.2 Safety Instructions for Sampling .............................................................................................. 14

1.3.3 Working and Personal Protection (Safety) Equipment ........................................................... 14

1.3.4 Use, Storage and Disposal of Reagents and Chemicals ........................................................... 14

1.3.5 Use, Storage and Disposal of the Equipment .......................................................................... 14

1.4 Environmental Protection ............................................................................................................... 14

1.5 Methods .......................................................................................................................................... 15

1.5.1 Prior to Sample Collection ....................................................................................................... 15

1.5.2 Water Sampling for Phytoplankton ......................................................................................... 16

1.5.3 Quantitative Analysis of Phytoplankton – general overview .................................................. 20

1.5.4 The Equipment used in Counting Phytoplankton .................................................................... 21

1.5.5 Sample preparation ................................................................................................................. 23

1.5.6 Taxonomy and Nomenclature ................................................................................................. 25

1.5.7 Counting Procedure ................................................................................................................. 26

1.6 Algal Objects and Counting Units .................................................................................................... 29

1.7 Identification and Coding ................................................................................................................ 29

1.8 Quality Assurance and Quality Control (QA/QC) ............................................................................. 31

1.9 References 1-Phytoplankton ........................................................................................................... 36

2. Zooplankton ............................................................................................................................................. 39





2.3.2 Safety Instructions for Sampling .............................................................................................. 42

2.3.3 Working and Personal Protection............................................................................................ 42

2.3.4 Use, storage and disposal of reagents and chemicals ............................................................. 43

2.3.5 Use, storage and disposal of equipment ................................................................................. 43


2.4 Environmental Indications ............................................................................................................... 43

2.5 Sampling Zooplankton ..................................................................................................................... 43

2.5.1 Prior to sample collection ........................................................................................................ 43

2.5.2 Required Strategies ................................................................................................................. 43

2.5.3 Preparation and calibration of sampling equipment .............................................................. 44

2.5.4 Sampling Equipment ................................................................................................................ 45

2.5.5 Sampling Design for Shallow and Deep Mesocosms ............................................................... 51

2.5.6 Best practice advice on preservation and storage .................................................................. 54

2.6 Quantitative Analysis of Zooplankton ............................................................................................. 55

2.6.1 Sample Preparation ................................................................................................................. 55

2.6.2 Counting Procedure (Enumeration) ........................................................................................ 58

2.6.3 Best Practice Advice on subsampling and counting ................................................................ 60

2.6.4 Taxonomy and nomenclature.................................................................................................. 61

2.7 Estimating the biomass ................................................................................................................... 61

2.7.1 Crustacean Zooplankton .......................................................................................................... 62

2.7.2 Rotifers .................................................................................................................................... 63

2.7.3 Estimating biomass: other methods in literature .................................................................... 65

2.7.4 Size Distribution ....................................................................................................................... 66

2.7.5 Size Diversity ............................................................................................................................ 66

2.7.6 Normalised biomass-size spectrum (NSS) ............................................................................... 66

2.8 Quality assurance and quality control ............................................................................................. 66

2.9 References 2 - Zooplankton ............................................................................................................. 68

3. Microbial Plankton .................................................................................................................................. 74



3.3 Health and Safety – Safe Disposal ................................................................................................... 76

3.4 Sample Integrity – Sampling – Best Practices .................................................................................. 77

3.4.1 Methods................................................................................................................................... 77

3.4.2 Processes ................................................................................................................................. 81

3.5 References 3 – Microbial Plankton .................................................................................................. 83

4. Periphyton ............................................................................................................................................... 85

4.1 Definition and Terms ....................................................................................................................... 85


4.3 Health and safety regulation ........................................................................................................... 85

4.4 Environmental Indications ............................................................................................................... 85

4.5 Methods .......................................................................................................................................... 86


4.5.1 Prior to sample collection ........................................................................................................ 86

4.5.2 Periphyton collection with artificial substrate (2) ................................................................... 87

4.6 Quality Assurance and Quality control ............................................................................................ 90

4.7 References 4 - Periphyton ............................................................................................................... 91

5. Water Chemistry ...................................................................................................................................... 92




5.2.2 Safety Instructions ................................................................................................................... 94

5.2.3 Working and Personal Protection (Safety) Equipment ........................................................... 94

5.2.4 Use, Storage and Disposal of Reagents and Chemicals ........................................................... 94

5.2.5 Use, Storage and Disposal of the Equipment .......................................................................... 95

5.3 Environment Indications ................................................................................................................. 95

5.4 Sampling .......................................................................................................................................... 95

5.4.1 Introduction ............................................................................................................................. 95

5.4.2 Sampling Strategy/ Sampling Plan ........................................................................................... 96

5.4.3 Equipment for Sampling .......................................................................................................... 97

5.4.4 Techniques for representative sampling ................................................................................. 98

5.4.5 Quality Assurance Considerations ........................................................................................... 99

5.5 Filtration ........................................................................................................................................ 100

5.5.1 Introduction ........................................................................................................................... 100

5.5.2 Vacuum Filtration .................................................................................................................. 100

5.5.3 Pressure Filtration ................................................................................................................. 101

5.5.4 Filter Types ............................................................................................................................ 101

5.5.5 Cleaning procedure for glass fibre filters .............................................................................. 102

5.5.6 Cleaning procedure for synthetic membrane filters ............................................................. 103

5.5.7 Volume for filtration .............................................................................................................. 103

5.6 Sample Storage .............................................................................................................................. 103

5.7 Auxiliary measurements ................................................................................................................ 105

5.7.1 Water transparency (aka Secchi depth) ................................................................................ 105

5.7.2 Light measurements .............................................................................................................. 105

5.7.3 Standard physical parameters (Temperature, Oxygen concentration, Conductivity) .......... 106

5.8 Methods applied by Project Partners ............................................................................................ 107

5.8.1 Aarhus University (AU) .......................................................................................................... 107

5.8.2 Centro de Biodiversidade e Recursos Genéticos – Universidade de Évora (CIBIO)............... 108

5.8.3 MARine Biodiversity, Conservation and Exploitation (CNRS-MARBEC) ................................ 110


5.8.4 Ecole Normale Superieure (ENS) ........................................................................................... 112

5.8.5 GEOMAR Helmholtz Centre for Ocean Research Kiel (GEOMAR) ......................................... 114

Hellenic Center for Marine Research (HCMR) ....................................................................................... 118

5.8.6 Ludwig-Maximilians-Universität Munich (LMU) .................................................................... 121

5.8.7 Middle East Technical University (METU) ............................................................................. 123

5.8.8 Netherlands Institute of Ecology (NIOO) ............................................................................... 125

5.8.9 Umweltbundesamt (UBA) ...................................................................................................... 126

5.8.10 University of Helsinki, Tvärminne Zoological Station (UH) .................................................... 132

5.8.11 University of Bergen (UIB) ..................................................................................................... 134

5.8.12 Umea Marine Science Center (UMF) ..................................................................................... 137

5.8.13 WasserCluster Lunz (WCL) ..................................................................................................... 140

5.9 References 5 – Water Chemistry ................................................................................................... 142

6. High Frequency Monitoring ................................................................................................................... 143

6.1 Definitions and Terms.................................................................................................................... 143

6.2 Equipment and supplies ................................................................................................................ 144

6.3 Health and safety indications ........................................................................................................ 144

6.4 Field Parameters and Sensors ....................................................................................................... 145

6.4.1 Temperature .......................................................................................................................... 145

6.4.2 pH........................................................................................................................................... 146

6.4.3 Dissolved Oxygen (concentration and saturation) ................................................................ 146

6.4.4 Conductivity ........................................................................................................................... 147

6.4.5 Underwater Light (PAR) ......................................................................................................... 149

6.4.6 Turbidity................................................................................................................................. 150

6.4.7 Depth ..................................................................................................................................... 152

6.4.8 Chl-a (including fluorescence per algal groups and total) ..................................................... 152

6.4.9 Nitrogen (Nitrate) .................................................................................................................. 153

6.4.10 Multi-probes (Sensors, profilers) ........................................................................................... 154

6.5 Maintenance of the station and the equipment ........................................................................... 155

6.5.1 Regular Sensor Inspection at the Field .................................................................................. 156

6.5.2 Best Practice Advice on Sensor-specific Field Cleaning and Calibration ............................... 159

6.6 Overcoming Drift ........................................................................................................................... 169

6.6.1 Common Causes .................................................................................................................... 169

6.6.2 Elimination strategies ............................................................................................................ 169

6.7 Troubleshooting of sensors and record-keeping equipment ........................................................ 170

6.8 Quality Assurance and Quality Control (QA/QC) of high frequency measurement ...................... 171

6.9 References 6 – High Frequency Measurements ............................................................................ 173


7. Data Quality Assurance and Quality Control ......................................................................................... 175

7.1 Definitions and terms .................................................................................................................... 175

7.2 Cross reference .............................................................................................................................. 175

7.3 Health and safety regulation ......................................................................................................... 175

7.4 Environmental indications ............................................................................................................. 176

7.5 Quality Assurance and Quality control Workflow ......................................................................... 176

7.5.1 Quality assurance of raw data collection .............................................................................. 177

7.5.2 Quality Control ...................................................................................................................... 177

7.5.3 Aggregated, summarized data............................................................................................... 183

7.6 References 7 – QA &QC ................................................................................................................. 184

III. Appendix ............................................................................................................................................ 185

Appendix I: Phytoplankton ........................................................................................................................ 185

Appendix I-B .......................................................................................................................................... 187

Appendix II: Zooplankton .......................................................................................................................... 189

Appendix II-A ......................................................................................................................................... 189

Appendix II-B ......................................................................................................................................... 191

Appendix II-C ......................................................................................................................................... 192

Appendix II-D ......................................................................................................................................... 194

Appendix II-E .......................................................................................................................................... 196

Appendix III: Microbial Plankton ............................................................................................................... 197

Appendix IV: Periphyton ............................................................................................................................ 197

Appendix V: Water Chemistry ................................................................................................................... 197

Appendix VI: High Frequency Measurements ........................................................................................... 198

Appendix VI-A: General Procedures for Calibration of Field Thermometers ........................................ 198

Appendix VI-B: General Procedures for Calibration of pH Sensors ....................................................... 200

Appendix VI-C: General Procedures for Calibration of DO Sensors ...................................................... 201

Appendix VI-D: General Procedures for Calibration of Conductivity Sensors ....................................... 203

Appendix VI-E: General Procedures for Calibration of in situ Fluorescent Sensors .............................. 204

Appendix VI-F: General procedures for 3-points calibration of ammonium and nitrate sensors ......... 204

Appendix VII QA &QC ................................................................................................................................ 205

Co-funded by the European Union D4.1 Standard Operating Procedures| I Executive summary

AQUACOSM – INFRA-01-2016-2017- N. 731065 of 205

I. Executive summary

This Standard Operating Procedure (SOP) describes methods for sampling and analysis of phytoplankton,

zooplankton, Periphyton, Water Chemistry, High Frequency Measurements, Data Quality Assurance and

Quality Control and Microbial Plankton from mesocosm experiments carried out in all aquatic environments

(fresh and marine waters). Additionally, it gathers best practice advice. This SOP is based on EU Water

Framework Directive and other related documents. It is designed to be compliant with this EU Directive

(2000/06/EC) [1]. Use of this SOP will ensure consistency and compliance in collecting and processing diverse

data from mesocosm experiments across the AQUACOSM community, in Europe and beyond.

II. Cross References

The SOPs that will be provided by AQUACOSM will be listed here in the following versions when the different

SOPs are completed.

The SOPs that will be provided by AQUACOSM will be for:

1. Phytoplankton (this SOP)

2. Zooplankton (Deliverable 4.1.2)

3. Microbial Plankton (Deliverable 4.1.3

4. Periphyton (Phytobenthos) (Deliverable 4.1.4)

5. Water Chemistry (Physical and Chemical Elements of Water) (Deliverable 4.1.5)

6. High-Frequency Data Collection (Deliverable 4.1.6)

7. QA/QC (Deliverable 4.1.7)

A general description for water sampling will be covered under the Water Chemistry SOP.

III. Dissemination activities related to the Deliverable

The SOPs will be made available to all users of TA in AQUACOSM, and will also be publicly available for any

user through the AQUACOSM webpage (https://www.aquacosm.eu/project-information/deliverables/)

https://www.aquacosm.eu/project-information/deliverables/

Co-funded by the European Union D4.1 Standard Operating Procedures| 1 Phytoplankton


1. Phytoplankton

1.1 Definitions and Terms

Biomass ● Total living organic material in a system or biological unit (e.g., taxon,

functional group) [2]

● Material composed of living organisms [3]

Biovolume Total cell volume of a taxon or a functional group per volume (e.g., L-1 water

sample) [2]

Cell volume The total volume of a single phytoplankton cell [2]

Epilimnion Upper water layer in stratified lakes that clearly differs from deep water layers

regarding water temperature. The depth of the epilimnion is defined as the

warmed uppermost and mixed water layer, with a relatively homogeneous

distribution of the water temperature during the summer stagnation period. It

lies above the metalimnion (thermocline), which is the horizon of the largest

vertical density change caused by differences in water temperature [2]

Euphotic zone The zone within a water body where photosynthetic production is possible

(gross primary production > respiration). It roughly corresponds to 2-2.5-times

of the transparency (Secchi depth). [2]

Metalimnion The transitional zone between the warm epilimnion and the cooler

hypolimnion. It corresponds to the lake zone with the largest density change

(thermocline; usually following the water-temperature gradient). According to

the original definition of BIRGE (1897), the metalimnion corresponds to the

zone where the change of water temperature within 1 m is at least 1 °C. [2]

Hypolimnion A stratum of cool water below the thermocline in stratified lakes.

Pelagic zone Open water zone of a lake, not close to the bottom or near the shore; in

contrast to the benthic or bottom zone of a lake [2]

Phytoplankton Community of free-floating, predominantly photosynthetic protists and

cyanobacteria in aquatic systems. (in limnological analysis commonly excluding

ciliates1). [2]

Secchi depth Maximal depth at which a b/w segmented disk (after Angelo Secchi) is just

visible from above the water surface. Measuring the Secchi depth (or

1 Ciliates with endosymbiotic algae, such as Mesodinium rubrum, are often abundant in oligotrophic waters and belong functionally to phytoplankton



transparency) allows determination of the turbidity of standing or running

water after ON EN ISO 7027 [2]

Taxon A phylogenetic unit (species, genus). Should be in accordance with current

phylogenetic classification

(algaebase.org as reference).

Thermocline See Metalimnion [2]

Trophic state The nutrient level of an aquatic system. Determines the biomass and primary

production of the community [2]

Utermöhl

technique

Technique for the quantification of phytoplankton through the sedimentation

of a preserved phytoplankton sample; specifically, determination and counting

of the organisms using an inverted microscope, calculation of the individual

biovolumes, projection of the counted organisms to a volume unit and

calculation of the total biovolume as described by Utermöhl (1958) with

modifications [2]

1.2 Materials and Reagents

Table 1-1: The materials and reagents used in the analysis of phytoplankton

Name and concentration Composition Storage

Lugol’s Solution Iodine based fixative most commonly

used for analysis of phytoplankton using

the Utermöhl technique. See Annex I for

detailed information about neutral, acidic

and alkaline Lugol’s solution.

Fume

Cabinet/Hood

Formaldehyde solution Fixative to be used for phytoplankton

analysis on the epiflorescent microscope.

See Annex I for detailed information about

preparing the formaldehyde solution.

Fume

Cabinet/Hood

http://www.algaebase.org/



1.3 Health and Safety Indications

1.3.1 General Information

In this section, general guidance on the protection of health and safety while sampling, analysing and

counting phytoplankton from mesocosm experiments will be provided to minimize the risk of health impacts,

injuries and maximize safety. The users of this SOP are expected to be familiar with the Good Laboratory

Practice (GLP) of World Health Organization (WHO) [4] and Principles on GLP of Organisation for Economic

Co-operation and Development (OECD) [5]. Health and Safety Instructions of the mesocosm facility, if there

are any, shall be followed properly to protect the people from hazardous substances and the harmful effects

of them.

According to preventive employment protection measures to avoid accidents and occupational diseases (on-

site or in the laboratory), the work should be practiced consistently with national and EU regulations (see the

OSH Framework Directive 89/391/EEC, [6]). Other regulations and guidelines can be found in the EU – OSHA

website (European directives on safety and health at work [7]). All necessary safety and protective measures

shall be taken by the users of this SOP, and the scientist-in-charge shall ensure that those measures comply

with the legal requirements.

The table below summarizes the hazards and risks and the measures for preventing them associated with

the laboratory studies carried out on phytoplankton:

Table 1-2: Hazards and risks associated with laboratory work

Occupations at

risk

Hazards/Risks Preventive Measures

Laboratories o Risk of inhaling

chemicals that are

used for phytoplankton

sample fixing

o Accidental spills

o Use of fume cabinets/hood

o Safe handling and transport of samples

o Appropriate personal protection (protective

gloves) and hygiene measures

o Samples are stored in glass bottles to

minimize diffusion of the fixative


o Use of lidded cool boxes to deliver samples

from field to laboratory

o Use of lidded plastic boxes to store the sample

bottles

http://www.who.int/tdr/publications/documents/glp-handbook.pdf



https://ntp.niehs.nih.gov/iccvam/suppdocs/feddocs/oecd/oecd_glpcm.pdf

http://eur-lex.europa.eu/legal-content/EN/TXT/PDF/?uri=CELEX:31989L0391&from=EN

https://osha.europa.eu/en/safety-and-health-legislation/european-directives



1.3.2 Safety Instructions for Sampling

Please see the Water Chemistry SOP (section 5, page 92) for safety instructions for sampling.

1.3.3 Working and Personal Protection (Safety) Equipment

Please see the Water Chemistry SOP (section 5, page 92) working, and personal protection equipment

suggested for use in water sampling.

1.3.4 Use, Storage and Disposal of Reagents and Chemicals

The user of this SOP needs to visit the Safety Data Sheets (SDSs), which is provided by the manufacturer for

any chemicals with the necessary information for protection before using, storing and disposing of the

reagents as well as other chemicals. Only experienced personnel should be responsible for the use of

reagents, preservatives, and chemicals in a way compatible with the laboratory rules if available.

It is important to remember that use of formaldehyde as a fixative can result in “the generation of

bis(chloromethyl)ether” - which is a potential human carcinogen [8]. Also, formaldehyde is an irritant and

can enter the body via inhalation and damage the tissues seriously. For more information on formaldehyde,

see the substance information ([9]) provided by the European Chemicals Agency (ECHA).

1.3.5 Use, Storage and Disposal of the Equipment

If phytoplankton sampling from the waters is to be carried out with a single piece of equipment, this

equipment needs to be cleaned in between two sampling events. [10]. The storage of equipment should be

compatible with the operating manual/instructions for use. Cleaning, disinfection, and storage of equipment

shall be taken care under the risk of parasites, diseases, foreign species, and pathogens.

1.4 Environmental Protection

A plan for the disposal of mesocosm waste needs to be prepared before the experiments. The plan must be

in competence with the EU Waste Legislation ([11]) and The List of Hazardous Wastes ([12]) provided by the

European Commission. The Safety Data Sheets (SDSs) need to be revisited for the disposal of reagents and

chemicals prior to waste disposal.

https://echa.europa.eu/substance-information/-/substanceinfo/100.000.002

http://ec.europa.eu/environment/waste/legislation/a.htm

http://eur-lex.europa.eu/legal-content/EN/TXT/PDF/?uri=CELEX:32000D0532&from=EN



1.5 Methods

1.5.1 Prior to Sample Collection

1.5.1.1 Introduction

The AQUACOSM Standard Operating Procedure (SOP) on qualitative and quantitative sampling, analysis and

counting of phytoplankton needs to be reviewed by the research team before initiating the sampling. The

sampling method to be employed, sample type and volume, equipment and supplies needed for sample

collection all shall be identified prior to the start of a sampling event.

1.5.1.2 The List of Required Equipment

The sampling equipment and supplies shall be provided by the AQUACOSM host facility. The following is a

list of equipment and supplies that can be used for recordkeeping. The list of equipment needed during the

sampling process is given in Section 1.5.2.1 and the list of equipment required for counting in Section 1.5.4.

The supplies for recordkeeping:

✓ Overview of mesocosm labels & treatments

✓ If mesocosms are stratified or contain macrophytes, the outline of sampling design (strata to be

sampled)

✓ Field sheets (Metadata Sheets) (Field protocol; DQ from [2])

✓ Markers and pens/pencils

✓ Camera

✓ Other relevant paperwork

✓ Notebook for contemporaneous notes if required

1.5.1.3 Calibration

The equipment should be checked. For sampler used in sampling and equipment for counting of

phytoplankton, the following methodology, as provided by the EU FP7 226273 (WISER), documentation, shall

be followed for calibration:

● Each counting chamber should be “marked with a unique mark or number and a note made of the

counting chamber area. This is calculated by measuring the cover slip aperture (rather than the chamber

itself) using either a Vernier gauge or the microscope stage Vernier if one is present. The mean of 5

diameters should be taken, and the area of the chamber calculated using the formula π r2” [13]. In

addition, the chamber volumes should be measured accurately by weighing the chamber (counting

chamber with cover slide + column of a determined volume + thick glass cover) and lid whilst empty, then

fill with distilled water and re-weigh (e.g., 5 mL chambers can range from 4.7-5.2 mL). The weight in

grams is equivalent to volume in mL. As a best practice advice, the calculation of volume should be



repeated three times, and the average of the three should be recorded. Both the diameters and the

chamber volume should be recorded in a log book.

● The “eyepiece/graticule and objective combinations should be calibrated with a stage micrometer (e.g.,

100μm x 10μm divisions) and the dimensions and areas of counting fields, transects and the whole

chamber area should be calculated for each of the magnifications used” [13].

1.5.2 Water Sampling for Phytoplankton

1.5.2.1 Equipment for Sampling

(Some of the equipment below will be moved to the Water Chemistry SOP when it is completed)

Sampling of phytoplankton is commonly carried out simultaneously with sampling for parameters of water

chemistry and chlorophyll-a. For general water sampling, check the SOP for Water Chemistry. The items

listed below are necessary for a water sample that is only used for phytoplankton analysis and does not

consider any further parameters.

The following specific pieces of equipment are suggested for use in phytoplankton sampling [2]:

✓ Appropriate water sampler, depending on the type of sampling (stratified or integrated), and (e.g.,

Schroder/Schindler/Ruttner sampler for single strata; tube sampler for depth integrated samples)

✓ Clean & rinsed sampling containers in the field; sample bottles for subsequent preservation and

storing of phytoplankton samples (Table 1-3 gives an overview of volumes)

✓ Supply of Lugol’s iodine for the preservation of samples

✓ A 10 or 20 µm plankton net can be used for concentrating phytoplankton. Net plankton samples can

be used to supplement the species list from the quantitative sampling.

Sampling containers and sample bottles for preservation [14]:

✓ Sampling containers: Depending on the sampling procedure (especially whether water is collected

for multiple parameters at one sampling event, or just for phytoplankton), water (the ‘sample’) from

the experimental units is either filled directly into bottle for fixation (‘sample bottle’), or a larger

amount of water is collected in a container (‘sampling container’), and the sample bottle is later filled

in the lab from the sampling container.

✓ Bottles for storing phytoplankton samples – ‘sample bottles’: The bottles should be selected

according to the purpose of the study and the type of sample. For Lugol’s fixation, transparent glass

bottles (50-300 mL, Table 1) should be preferably used, as iodine will diffuse through standard plastic

containers (e.g., PE, PP, PS). Transparent glass bottles should be preferred since it is easier to see if

the amount of Lugol’s is enough through a transparent bottle. Sample bottles should be stored dark

and cool. Keeping them in a fridge helps to preserve the material (temperature ca. +4 - +8 °C). It is

essential that the user of this SOP ensure sample bottles are stored and transported clean, ideally in



their original packaging. Used bottles can be re-used, if cleaned and rinsed properly (otherwise cells

sticking to the bottle walls might contaminate new samples).

✓ Best practice advice: Lugol's solution undergoes photo-degradation, it is recommended that samples

are stored in brown glass bottles.

✓ Best practice advice: “It is important that the bottle cap is securely tightened to avoid spillage of the

sample and evaporation of the preservative. Utermöhl (1958) ([15]) recommended that the bottle is

filled to 75-80% of its volume. This facilitates the homogenisation of the sample before dispensing

into the sedimentation chamber” [16].

Labelling:

✓ Sampling containers and sample bottles need to be appropriately labelled in the laboratory before

sampling. The labels on the sampling bottles need to be standardized and provide information on

the name of the experiment, sampling date, mesocosm ID, and possibly depth with appropriate

abbreviations of the treatments of the experiments. Labels can be either printed or written using a

permanent waterproof marker [14].

✓ As a best practice advice, a label template is provided in Annex II of this SOP.

1.5.2.2 Sampling

Representative sampling – samples need to be representative of the mesocosm unit of interest. In case other

parameters (esp. nutrients, chlorophyll-a) are sampled at the same event, it is mandatory that all parameters

are analysed from the same water sample (multiple sub-samples from one water sample).

A. Mixed mesocosms

In case mesocosms are mixed continuously, one water sample can be considered representative for the

entire mesocosm. Here, samples should be taken near the center of the mesocosm as stated in both [13] and

[17]. Care should be taken to avoid macrophytes, if present, while sampling. If it is unclear whether the whole

water volume is efficiently mixed, an initial series of samples can be taken in transects across the mesocosm

and along the vertical axis (see point 7.2.2 B below).

B. Stratified mesocosms

If mesocosms waters are stratified or partially mixed, the sampling procedure must be determined after

careful considerations on the water layer(s) to be sampled (e.g., sampling discrete water layers; sampling

multiple depths with subsequent pooling to one combined representative sample; utilisation of tube sampler

for integrated water layers). The absence of vertical temperature profiles indicates vertical mixing, but NOT

necessarily homogeneous distribution of motile organisms like (micro-) zooplankton. Heterogeneous

distribution of phytoplankton must also be assumed if mesocosms contain structuring elements (such as

macrophytes), and has not been proven to be (practically) homogeneous (see 7.2.2.A).



C. Composite and discrete sampling

According to WISER [13], representation of the whole phytoplankton community will be achieved, taking

vertically integrated samples from the euphotic zone in a thermally stratified mesocosm. Careful

consideration of the sampling design is also required in case the mesocosms are stratified and include an

aphotic zone (mesocosm depth > Zeu2). In this case, the integrated water sample would typically be taken

from the euphotic zone (Zeu).

For shallow mesocosms containing sediment and possibly macrophytes, a detailed sampling design for

collecting samples at multiple horizontal and vertical positions might be needed [17]:

“The entire water column, from the surface to approx. 5 cm above the sediment, is sampled from

three positions in each enclosure: 10, 30 and 60 cm from the enclosure wall. Two samples are

prepared: one to be used for chemical and phytoplankton analyses where water is sampled without

touching the plants – and one to be used for zooplankton analysis, where water is also sampled close

to the plants.” “The best way to sample from the entire water column is by using tube sampler which

sample from top to bottom. The diameter of the tube should not be too small to avoid zooplankton

escaping during sampling (> 6 cm). If it is not possible to use a core sampler, samples can be taken

with Ruttner water sampler from the surface (20 cm below the water surface), middle and the

bottom (20 cm above the sediment). The sample in the middle should be adjusted according to the

enclosure type and actual water depth” [16].

D. Recommended sample volume

Table 1-3 below gives a rough approximation about the required sample volume (total sample volume and

volume for sedimentation): Note that the phytoplankton sample volume needs to be large enough to allow

for preparation of multiple sedimentation samples (i.e., at least 3x the volume of the sedimentation chamber

to be used).

Table 1-3: Recommended sample volumes and volumes for sedimentation according to the trophic state of an aquatic system

Trophic state (TP µg L-1) Sample bottle volume (ml) Volume for sedimentation (ml)

Ultra-oligotroph (<<10 µg) 500 ≥100 (concentrate)

Oligo-mesotroph (10-25 µg) 150-250 25-50

Meso-eutroph (15 - 45 µg) 50-150 5-25

Eutroph (>= 50 µg) 10-30 < 5 ml (dilute)

2 Zeu corresponds to 2-2.5 x Secchi depth



1.5.2.3 Sample Fixation

Phytoplankton samples are usually fixed by Lugol’s solution (acidic or alkaline solution, see Annex 1). Lugol’s

iodine preserves the cells. In addition, iodine increases the specific density of cells and enhances

sedimentation.

“Use alkaline Lugol’s solution (using sodium acetate buffer) or acidic Lugol’s (which allows better

sedimentation of buoyant cyanobacteria) as a preservative to reach a final concentration of about

0.5% in the sample, i.e., about 8 drops per 100 mL (or 2.5 mL for a 500 mL flask). The final

concentration should give the sample a light brown/orange colour (whisky or cognac colour).

Depending on the type of sample, reaching the colour can take a higher number of drops – in acid

waters for instance” [13].

Lugol-preserved samples should be stored in the dark and temperature 4 to 8 degrees of Celcius until

analysis. Samples should be analysed within a few months. If longer-term storage is needed, the intensity of

coloration should be checked optically every 6 months to ensure sufficient concentration of the fixative (loss

of coloration indicates loss of fixative by diffusion). ✓ As a best practice advice, it is essential to leave an-airspace of approximately 1-2 cm below the lid

in the bottle. If the bottle is filled to the top, mixing (by shaking) of the sample for preparation of the

sediment chamber will be inefficient.

Fixation with formaldehyde (or glutaraldehyde) is used in combination with epifluorescence analysis (the

sample is then filtered on a dark membrane filter before microscopic analysis and stained by a DNA dye). The

advantage of this method is it allows a careful distinction of pigmented from non-pigmented cells. The proper

procedure for preparing a formaldehyde solution is provided in Annex 2 of this SOP. The recommended final

concentration of the fixative (formaldehyde or glutaraldehyde) is 2%.

✓ It is important that the Lugol’s and especially formaldehyde solution be stored and used in the Fume

Cabinets (Fume Hood) while using them for preserving samples. The material used in combination

with form-/glutaraldehyde should be labelled for this purpose. It is recommended to clean this

material using gloves. Country-specific regulations may apply esp. regarding the use of Formaldehyde

and Glutaraldehyde.



1.5.3 Quantitative Analysis of Phytoplankton – general overview

The workflow of the quantitative analysis of phytoplankton is summarized in Figure 1-1 below.

Figure 1-1: The Workflow of quantitative analysis of phytoplankton



1.5.4 The Equipment used in Counting Phytoplankton

The equipment that can be used in counting phytoplankton (adapted from WISER [16])

● “Sedimentation chambers of 5 to 50 ml capacity. Chambers should be approximately 25 mm in

diameter. Sedimentation chambers of ≥ 100 mL should be avoided especially for small cells

because of the high risk of uneven and non-quantitative sedimentation of small cells. Instead,

dilute or concentrate samples if densities are very high or low (the concentration is too high if

cells lay on top of each other; too low concentration is indicated if the required number of units

counted per sample cannot be achieved, see below)”. [16]

- Combined plate chambers: A tower is placed on top of a base plate (Figure 1-2.B-D). The

sample is filled into the chamber. After sufficient time has passed (Table 1-4), the tower

is removed.

- Tubular chamber: The tower is permanently fixed on the settling chamber. The sample

should be placed in humid chamber already for sedimentation to avoid the formation of

gas bubbles. (Figure 1-2.A). Tubular chambers allow sedimentation of limited volume (5-

10ml). Note that the height of chamber inc. tower must fit under the condenser unit of

the microscope.

Figure 1-2: Settling chambers. A: tybular chamber with a fixed tower, different volumes. The sample volume stays in the chamber during analysis. B-D, combined plate chambers. B, accessoirs. C, a chamber with the sample. D, Chamber after removing the supernatant. The tower with the sample volume is gently pushed to the side by the cover glass plate. Pictures are taken from https://www.hydrobios.de/ .

https://www.hydrobios.de/



Best Practice Advice: “The cleaning of sedimentation chambers is a critical part of the Utermöhl technique.

The chambers should be cleaned immediately after analysis to prevent salt precipitate formation. A soft

brush and general-purpose detergent should be used. To clean the chamber margin properly, a toothpick can

be used. Usually, it is sufficient to clean the chamber bottom without disassembling the bottom glass.

Sometimes, however, it is necessary to separate the bottom glass from the chamber, either to clean it or to

replace it. This is easily done by loosening the ring holding the bottom glass with the key. Care should be

taken as the bottom glasses are very delicate. Counting chambers should be checked regularly to ensure that

no organisms stick to the bottom glass. This can be achieved by filling the chambers with distilled water” [16].

● Inverted microscope with phase contrast (and/or Differential interference contrast) including:

- Standard resources for microscopy (slides, cover slips, pipettes, cleaning paper, wash bottle,

etc.)

- long working distance condenser with numerical aperture of >0.5

- 10x or 12.5x binocular eyepieces; preferably one with a square grid, and another with a cross-

hair graticule with scale bar (see Figure 1-3)

- 10x, 20x and 40x phase &/or Differential Interference Contrast (DIC) objectives

- ideally, the microscope should be fitted with a digital camera coupled to image analysis

system

- a mechanical stage

● Supply of deionized/degassed or membrane filtered water is recommended for topping up,

diluting and general cleaning” [13].

Figure 1-3: Examples of suitable eyepiece graticules (revised from [13])



1.5.5 Sample preparation

1.5.5.1 Acclimatization of sample and counting chambers

Samples and equipment involved in preparing Utermöhl samples should be acclimatized for 24

hours/overnight to room temperature. This is important in achieving random sedimentation of algal cells on

the bottom of the settling chamber [13].

1.5.5.2 Sample Mixing

Before sub-sampling, the sample should be mixed following the methodology provided by the WISER:

“Just before taking a sub-sample to fill the sedimentation chamber, the sample is manually

thoroughly mixed using a combination of alternating horizontal rolling and vertical tumbling (turning

upside down) of the sample bottle for around 2 minutes. These actions should be gentle and not

involve any vigorous shaking or vortex formation” [13].

1.5.5.3 Sub-Sample Volume

The exact volume of sample used to fill the chamber depends on the phytoplankton density (see Table 1-3).

Some options are available for dealing with varying densities of phytoplankton (adopted from [13]):

● Use a sedimentation chamber of an appropriate size depending on how abundant the phytoplankton

cells are (chlorophyll-a and/or nutrient concentration may serve as guidance, see Table 1-3). If there

is enough sample water to be used for several sedimentations, it is recommended to use a couple of

different sedimentation chambers with different volumes to prepare a sample for microscopy. After

sufficient sedimentation time, samples are checked with a microscope and a sample in which

phytoplankton density is the most appropriate and evenly distributed is selected for the analysis.

● It is recommended to use settling chambers between 5 ml and 50 ml. Use of chambers with smaller

or larger volumes is likely to result in uneven distribution of small cells.

● For very low densities, a pre-concentration step may be necessary. Let sample settle in a measuring

cylinder - usually, 250 mL is sufficient. Leave for 3 days and then draw off top water leaving 25 mL at

the bottom of the cylinder (i.e., x10 concentration). If needed this can be repeated with up to 4 250

mL cylinders, and the 4 lots of 25 mL then poured into a 100 mL measuring cylinder for a second pre-

concentration to 10 mL (i.e., x100 concentration) [18].

● In ultra-oligotrophic systems, large settling towers (>50 mL) can be employed for counting large

phytoplankton (microplankton, cells ≥20µm; large cells sediment faster than nano- and picoplankton,

hence the risk of uneven distribution is limited).

● For very dense samples, it is necessary to dilute the sample before adding it to the chamber. Add a

known volume of sub-sample to a measuring cylinder and top-up to a measured volume with Lugol’s

preserved water, thereby ensuring proper mixing of the sample with the added water. The water



used for dilution must be particle free. In freshwater analytics, deionized water can be used. In

marine mesocosms, seawater filtered by a 0.2 µm filter will be better as deionized water might cause

cells to disintegrate.

● As an alternative to dilution of samples, Palmer chambers can be used for enumeration of dominant

taxa in very dense samples (volume of a Palmer chamber is typically 100µL). Palmer chambers do not

necessarily warrant even distribution of cells. Ideally, the entire chamber is counted.

1.5.5.4 Proper Environment and Required Time for Sedimentation

The time required to ensure quantitative sedimentation of all particles including small phytoplankton

depends on the height of the settling chamber. A good recommendation is to allow for 4h of sedimentation

per centimeter of settling chamber height. Following the WISER guideline [13], one should allow “10 ml

chambers to settle for at least 12 hours, 25 ml chambers for at least 24 hours, and 50 ml chambers for at

least 48 hours. Note that too long a settling period (several days) increases the risk of disturbance and

formation of air bubbles.” Table 1-4 below gives an overview about settling time).

Best practice advice:

✓ Even distribution of particles requires the settling chamber to be placed on a horizontal surface.

Especially for larger settling chamber (20 mL and more), it is advisable to check the horizontal

orientation of the surface by a tubular level.

✓ The table used for preparing the settling chamber must not be vibrating or shaking. There should be

no other equipment on the same table during sedimentation that might cause vibrations (like a

vacuum pump). The table should not be placed on a hardwood floor which vibrates, i.e., when people

pass the room.

✓ Air bubbles inside the settling chamber disturb the vision and resolution when counting. To minimize

air bubbles, ensure the acclimation of material (see above) and store the settling chamber in a humid

environment (Figure 1-2). Such a space is easily created by placing the chamber next to a petri dish

filled with water and covering both items under a plastic bowl turned upside down. When employing

combined plate chambers, this procedure is relevant for the time after the settling tower has been

removed.

In addition, HELCOM COMBINE [19]; a manual for monitoring the marine waters, provided the minimum

amount of settling times; which are dependent on the height of the chamber as well as the type of

preservative used. The given settling time for specific chamber volumes and height when Lugol’s is used as

the preservative are tabularized below:



Table 1-4: Minimum amount of settling times for specific chamber volumes and height (adopted from [19]). The settling time depends mostly on the height of the settling water column, with ca. 4 h per cm.

Volume of Chamber

(cm3)

Height of Chamber

(cm)

Settling Time

(h)

2-3 0.5 – 1 4

10 2 12

25 5 24

50 10 48

1.5.6 Taxonomy and Nomenclature

The nomenclature and taxonomy should follow best practice. For general inquiries about phytoplankton

taxonomy and nomenclature, AlgaeBase [22] is a good reference for both freshwater and marine taxa. Also,

the WORMS database (http://www.marinespecies.org/) is currently the best reference for marine taxa (and

continuously synchronized with the AlgaeBase). For Baltic Sea phytoplankton, the taxa list of HELCOM

Phytoplankton Expert group (PEG) [21] should preferably be used. Additionally, IGB in Berlin is currently

elaborating a taxa list [20] to be used for lakes. Recommended literature for identification of freshwater,

brackish and marine taxa:

● For freshwater phytoplankton, an extensive overview of available literature is given in [13].

● For marine phytoplankton, UNESCO/IOC [16] suggested the review of the following literature for

identification: Horner (2002) [23], Tomas (1997) [24] and Throndsen and others (2007) [25].

● One can also see Santhanam and others (2019) [47] for taxonomic identification of marine

phytoplankton.

● For Baltic Sea phytoplankton, species lists are published on the webspace of the HELCOM

Phytoplankton Expert Group [21].

✓ Best practice advice: It is highly recommended to speak to local experts about the recommended

literature and check with them about species lists of the local habitats.

Taxonomic resolution

The observed taxa are identified to the highest possible taxonomic level. It is critical to remember that it is

better to correctly identify algae to lower taxonomic level than misidentify to a higher level (e.g., report

genus level if species cannot be identified unequivocally).

http://www.algaebase.org/

http://www.helcom.fi/helcom-at-work/projects/phytoplankton


http://www.igb-berlin.de/en/project/taxaliste-phytoplankton-o-217



Notes on the definition of phytoplankton

Phytoplankton is defined as the sum of suspended photosynthetic protists and cyanobacteria. However, the

most commonly used method for analysing phytoplankton (Utermohl method in combination with Lugol’s

solution; this SOP) does NOT allow to distinguish pigmented (photosynthetic) from non-pigmented cells.

Furthermore, the method is not suitable for quantitative analysis of ciliates in the sample (ciliates may be

auto- mixo- or heterotrophic). The analyst should be aware of the following issues:

● Auto- mixo- and heterotrophic protists: The recent appraisal of widespread mixotrophy in

phytoplankton [26] renders the traditional distinction of “autotrophic phytoplankton” and

“heterotrophic protozoa” inoperative.

● There is no consensus whether heterotrophic protists should be counted (e.g., heterotrophic

dinoflagellates such as Gymnodinium helveticum, Protoperidinium sp.). The identifier must be aware

that Lugol fixed samples do not allow distinguishing pigmented from non-pigmented cells (fixation

with formaldehyde in combination with epifluorescent microscopy is required, see 7.2, above).

Heterotrophic taxa should be flagged in the datasheet when identified as such.

● Especially in freshwater studies, ciliates are typically not counted within phytoplankton analysis.

Note, however, that many ciliates are mixotrophic (combine photosynthesis with phagotrophy).

● Many ciliates will not be retained quantitatively in Lugol’s fixed samples If ciliates are to be counted,

samples fixed with acidic Lugol’s solution can be prepared for enumeration of microzooplankton, inc.

ciliates [18]. If ciliates are a particular focus of a study, specific samples for ciliates should be prepared

using appropriate fixatives (like Bouin's fixative [27]).

● In the Baltic Sea phytoplankton monitoring, the endosymbiotic ciliate Mesodinium rubrum is

counted, since it contributes to primary production. Thus, M. rubrum should preferably be counted

also within the Baltic Sea mesocosm studies (to be able to compare the mesocosm results with the

long-term monitoring results, etc.).

1.5.7 Counting Procedure

An overview of the general principles for quantitative analysis of phytoplankton is provided by the WISER, as

follows:

“The quantitative analysis described here includes the identification, enumeration, and calculation

of biovolumes of Lugol’s iodine preserved water samples. The analysis should be carried out using

sedimentation chambers with an inverted microscope (Utermöhl technique).

The preserved sample is thoroughly mixed, and a sub-sample of known volume is placed in a

sedimentation chamber. When the cells have settled to the bottom of the chamber, they are counted

and identified using an inverted microscope.



The statistical reliability of the analysis depends upon the distribution of phytoplankton units/cells

within the sedimentation chamber and assumes that the cells are randomly distributed within the

chamber.

The counts for individual taxa are converted to phytoplankton biomass by using the cell/unit volume

of the count units. The volumes are based on measurements made during counting or on available

biovolume information for different taxa and size-classes.” [13]

The following instruction on the counting procedure is adopted from the corresponding WISER document

[13].

✓ It is useful to scan the sample at a variety of magnifications before the quantitative analysis is

undertaken and to compile a list of the most frequent taxa before beginning to count.

Counting at different magnifications (steps A, B, C below)

To obtain a reliable estimate of relevant size classes, the countings should be carried out at 2-3 different

size classes in the following manner (note that the effective magnification is the combined magnification of

objective and ocular):

• at low magnification (40x or 100x), a whole chamber count to pick up large taxa, followed by;

• 2 transect counts at an intermediate magnification (200x or 250x), to enumerate “intermediate-

sized” taxa (>20 μm) that are too small for the low-magnification count but too large to be reasonably

counted using fields of view at high magnification, followed by;

• A high magnification count (x400 or greater) using fields of view to pick up the small taxa. Aim to

count 50-100 fields of view (i.e., at least 400 units assuming the recommended sample

concentration).

More details are provided in the sections below.

1.5.7.1 Counting the whole chamber at low magnification for large taxa

Working at low magnification (x40 to x100) the whole chamber should be scanned in a series of horizontal

or vertical transects (Figure 1-3a), and the larger taxa (e.g., large dinoflagellates), large colonial or filamentous

forms (e.g., Microcystis, Fragilaria) are counted. A cross-hair graticule eyepiece, or similar, (Figure 1-3) should

be used if possible when counting the whole chamber. In horizontal transects, algae that lie between two

horizontal lines are counted as they pass the horizontal line; algal objects that cross the top line are included



while those crossing the bottom line are not and will be counted on the next transect (or vice versa; proceed

accordingly for vertical transects).

1.5.7.2 Counting diameter transects

Algal cells larger than approximately 20 μm (e.g., Cryptomonas) are counted at 200x – 250x magnification in

2 randomly chosen diameter transects of the counting chamber (Figure 1-3B). The cross-hair eyepiece and

method for counting algal objects described in the section above are used. The chamber is rotated between

transect to randomly chosen positions.

1.5.7.3 Counting randomly selected fields

Small algae, <20 μm (e.g., Rhodomonas, small centric diatoms), should be counted in 50-100 randomly

selected fields at x400 magnification (or greater) using a square, Whipple graticule, Miller Square or similar

in the ocular eyepiece or the field of view to delineating the counting area. The number of fields counted

should achieve a total count of approximately 400 phytoplankton units for the sample. Fields should be

selected in a stratified-random way following the same pattern as the full chamber counts (Figure 1-4A). The

counter must not look into the microscope when selecting a field – as this will result in non-random selection

of fields.

Figure 1-4: A: Counting method for the whole chamber; horizontal transects, B: Counting method for diameter transects, C: Exemplified rule for counting cells at the edge of the field

A tally of the number of fields counted is required as well as the counts of individual identified algal units

(cells, colonies or filaments).

When counting random fields, it is important to take a consistent approach deciding whether unicellular algal

objects lying across the grid lines are counted in or out. A simple rule should be adopted as described in the

CEN method (2006): unicellular algal cells crossing either the top or the left-hand side of the grid are not

counted while those crossing the bottom or right-hand side of the grid are counted (Figure 1-4C).



For filaments and colonies, only the cells or filament length that is inside the field of view should be counted.

Ideally, however, these larger taxa should usually be counted at lower magnification in transects or full

chamber count.

1.6 Algal Objects and Counting Units

● Dead cells and empty loricas should not be counted (e.g., Dinobryon, Cyclotella)

● Littoral or benthic taxa should be counted and flagged as such (especially if mesocosms contain

benthic compartment). In shallow systems, they can contribute a significant proportion of the

sample.

● Heterocysts and akinetes of filamentous cyanobacteria should be counted (with separate

measurements for biovolume estimates if present in large number).

● Unicellular picoplankton (<2 μm) should not be counted (unless using epifluorescence microscope).

Counting units are independent algal cells, chains, colonies or filaments/trichomes. One species or taxa may

be present in the sample as different counting units and may be counted at different magnifications.

For example, Microcystis colonies are counted in the whole-chamber or transect, but individual Microcystis

cells (which may be present if colonies are disintegrating) are counted in random fields. Similarly, Dinobryon

colonies may be counted in whole chamber or diameter transects, but single Dinobryon cells often need to

be counted in random fields. Chain-forming algae (e.g., Chaetoceros) can be counted as single cells or

colonies.

Counting units are assigned following practical considerations, E.g., some filamentous forms do not allow

distinction of individual cells. (e.g., counting Planktothrix sp. in units of 100µm filament length).

Species with a high variation of size can be counted in size-classes (e.g., Cryptomonadales <16 μm, 16-26 μm,

>26 μm).

For analyses concerning Baltic Sea phytoplankton samples, counting units as given in the HELCOM PEG taxa

and biovolume list [21] should preferably be used.

1.7 Identification and Coding

Species are coded as presented in the WISER_REBECCA taxa list [28] for phytoplankton available on the

WISER intranet. The present updated taxa list is also included in the counter spreadsheet.

● Calculation of phytoplankton biovolume

Biovolumes must be measured for all taxa, and it is done by assigning simple geometric shapes to each cell,

filament or colony, measuring the appropriate dimensions and inputting these into formulae to calculate the

cell volume. To obtain good biovolume estimates, species-specific biovolume should be estimated for all



http://www.freshwaterecology.info/TaxaDB_phpSearch.php



abundant taxa. For rare taxa, biovolume from previous analyses (same system) or even from published

resources can be used (e.g. [29],[30])

Hillebrand et al. [31] provide a comprehensive overview of formula for calculating biovolumes in

phytoplankton. Most of the formula is available in the WISER excel spreadsheet for data entry (see Data

entry, below).

For analyses concerning Baltic Sea phytoplankton samples, taxon and size class-specific biovolumes given in

the HELCOM PEG taxa and biovolume list [21] should preferably be used.

This SOP comes with an excel datasheet for entry of count data (adapted from [13]; see Data entry, below).

The sheet contains species lists and formula for the biovolume calculation for of each taxon. The cell

dimensions that need to be measured depend on the underlying geometrical shape (see points listed below).

Measurements of the required cell dimensions (length, width, diameter) are made at an appropriate

magnification using a calibrated ocular eyepiece, for instance, a Whipple Graticule. The eyepiece is rotated

so that the scale is put over the required cell dimension and the measurement made by taking the ocular

measurement and multiplying by the calibration factor for that magnification and eyepiece combination. The

measurements can also be made by image analysis. Per taxon, ca. 20 randomly chosen cells should be

measured.

● Biovolume estimation of unicellular taxa

It is important to measure the linear dimensions of some individual units of all taxa observed in the sample.

For taxa of more variable size (e.g., centric diatoms), and taxa that contribute significantly to the total

biovolume (e.g.,>5% of biovolume), at least 10 individuals should be measured.

For some species with external skeletons much larger than cell contents, e.g., Dinobryon, Urosolenia, the

dimensions of the plasma/organic cell contents should be measured, not the external skeleton dimensions.

● Biovolume estimation of filamentous taxa

For filamentous taxa, the average biovolume can be approximated by estimating the number of cells per

filament multiplying by the mean biovolume of one cell. Alternatively, it is possible to use the mean

dimensions of filaments to calculate the biovolume of one filament multiplying by the number of filaments.

● Biovolume estimation of colonial taxa

For colonial taxa, count or estimate cell numbers and multiply them by mean cell dimensions (Often, single

measures of dimensions are needed). Using colony/coenobium measurements – measure colony width and

depth e.g., Pediastrum – with colony depth approximated as an individual cell diameter.

For dense colonies, it may be difficult to estimate cell numbers. Some labs recommend estimating biovolume

for e.g. Microcystis by applying a reduction factor on the biovolume of a colony (RF = reduction factor = 0.4;

d = diameter of spherical colony; biovolume = RF x 𝜋/6 x d3) [32]. Similar to cellular biovolume, density of




colonies is not a constant factor but varies, i.e., with age of colony. For a given sample, such a factor must be

validated by careful microscopic inspection of a number of colonies.

● Note on counting effort, taxon richness & diversity estimates

If diversity parameters (taxon richness, evenness) are of interest, it is critical to maintaining a constant

counting effort among samples. A good standard is to count a minimum of 500 counting units per sample.

Rarefaction analysis (as described in [33]) should be performed to check for sampling bias regarding the

parameter of interest. Constant counting effort also implies to balance detection limit relative to the

biovolume of taxa (large taxa contribute over-proportionally to the total biovolume per sample). That is, the

counting effort should be similar for different magnifications. As a general recommendation, the time spent

on counting at different magnification should be in a similar range.

● Data entry

The counting can be carried out using different programs or spreadsheets. A WISER excel spreadsheet that

was prepared during the WISER project is recommended for freshwater taxa

(https://www.aquacosm.eu/project-information/deliverables/). The spreadsheet contains the whole

WISER_REBECCA taxa list [28] and provides biovolume formulae for many of the common taxa (the overview

of formula can also be used as a general reference for biovolume calculation). It also allows the raw data to

be summarized. All required details must be input into the counting spreadsheet according to the

accompanying instructions.

Data to be entered will include information on the sample ‘site’ (i.e., mesocosm ID) and date of collection,

date of analysis, the person who carried out the count, information on the chamber and counting areas and

the volume of sample used. For each taxon found, the number of units counted, the number of fields of view

(or equivalent for whole chamber or diameter transects) in which it was counted and mean dimensions will

be recorded for a given taxon. For taxa which are counted in different counting units (e.g., individual cells

and filaments/colonies), it is important to fill in one row for cells counted and the other for filaments or

colonies. For filaments and colonies, an estimate of the numbers of cells is also usually required to calculate

biovolume/mL. Cells/mL and biovolume/mL for each taxon are automatically calculated once the count and

mean dimensions are entered.

1.8 Quality Assurance and Quality Control (QA/QC)

According to [14], systematic or random errors can occur during the sampling operations. Systematic errors

are generally due to the “poor sampling practices or equipment design failures” which are usually constant.

On the other hand, random errors are generally unavoidable or unpredictable. One shall follow an effective

Quality Assurance/Quality Control (QA/QC) strategy during the experiment to identify, quantify and control

the errors. Standardization of sampling and analysis methods, taking replicate samples and analyses, or

following a laboratory accreditation scheme are some examples of QA practices that can be followed during

http://www.freshwaterecology.info/TaxaDB_phpSearch.php



the experiment. Quality Assurance (QA) strategies are defined by the scientist-in-charge to ensure that the

sample data meets Data Quality Objectives (DQO), which shall be defined before sampling. More information

on DQOs can be found in the separate SOP on Data Quality Management (Reference will be provided for

DQM SOP). On the other hand, Quality Control (QC) is “the system of guidelines, procedures and practices

designed to regulate and control the quality of products and services, ensuring they meet pre-established

performance criteria and standards.” The QC practices that can be followed are as follows: taking “sample

blanks, replicates, splits,” having and following “equipment calibration standards,” determination of “sample

container size, quality, use, and preservative amount” prior to sampling.

The WISER project results highlighted the information below for QA of sampling and counting of

phytoplankton:

“1) Details of microscopes, chambers (individually identified and calibrated) and calibration of all

ocular/objective combinations should be recorded in a notebook and kept for reference. If fixed

volume pipettes are used, these should be calibrated annually.

2) Checks for a random distribution of sample should be done visually at low magnification for each

sample. Some simple checks include:

Comparing the number of observations in:

(a) half a chamber with the other half

(b) comparing counts in the 1st transect with the 2nd transect

(c) comparing counts in the first 20 field of view with the next 20 fields.

A more detailed check using simple Chi-squared test should be done if a sample does not appear to

be randomly sedimented or 1 sample every 3 months or so” [13].

Also, the following were also suggested as a best practice advice by Brierley and colleagues (2007):

● If there is an aim for accreditation of phytoplankton analysis, ring-tests can be undertaken with

a staged approach:

1) “Determining mainly counting errors – group of analysts to count a limited number of

named taxa (1 to 3) or latex particles/pollen grains in set fields – can be done using

photographs or videos

2) Repeat transect or field counts by 2 or more analysts on real sample to check

identification and counting errors.

3) Full count comparisons

● Regular workshops should be held (3 - 4 times per annum) to carry out identification and ring

tests, possibly combined with 1⁄2-1 day taught workshop on difficult groups [34].

Moreover, UNESCO/IOC [16] determined a quality assurance procedure to follow when using the Utermöhl

technique. According to the document, the quality of the results is dependent on the skill of the analyst and,



to ensure high quality results, one shall validate all steps of the method. The steps in the Utermöhl method

to validate are determined in the document as follows:

i. homogenization of sample

ii. sedimentation/sinking

iii. distribution on chamber bottom

iv. repeatability and reproducibility” [16].

Rott and others (2007) [35] suggested the use of both “intra-laboratory standardization and inter-(between)

laboratory calibration (ring-test)” for quality assurance of the quantitative analysis of phytoplankton. In

addition, the use of the following results of his study as a pathway to assess counting precision:

● “Repeated subsample counts (pairs of transects, random fields, etc.) evaluated separately can

be simple and quick approaches for intra-laboratory scatter check, and can be helpful in

identifying quality targets, which can be independent from the actual subsampling methods

● Minimum counting thresholds for unicellular taxa were found to be attained by counts of less

than 50 in spite of fundamentally different approaches

● Although conventional size measurements and protocols (e.g. [36]; [37]) can be precise enough,

traditional shape analysis can be improved. Microscopic observations combined with novel

electronic methods ([38]) and flexible 3D electronic models ([39]) are recommended for future”

[35].


✓ The mesocosm facilities can work in a network. ‘Nordic Periphyton & Phytoplankton Group’ (NPPG), is

an example of such a network, which organizes annual meetings and practices inter-calibration [40]. The

group is made up of specialized algologists.

✓ In addition to NPPG, Nordic Microalgae [41] for the mesocosms in the Nordic Region can be examined

for information about microalgae and related organisms.

✓ For the Baltic Region, HELCOM Phytoplankton Expert Group (HELCOM PEG) project website [42] can be

visited to receive information especially on Baltic Sea phytoplankton species and QA procedures

followed.

✓ For sampling marine phytoplankton, PLANKTON*NET [43] data provider at the Alfred Wegener Institute

for Polar and Marine Research (AWI) is suggested as an open access repository for plankton-related

information.

For detailed QA methodology provided for sampling, analysis, and counting of phytoplankton, please visit

CEN - EN 15204 [44], Rott et al. [35] and WISER project [13].



According to the ‘Common Implementation Strategy for the Water Framework Directive (2000/60/EC)’, the

inter-calibration exercises in between laboratories will provide a “continuous quality assurance system,” by

ensuring the results meet targeted levels [45]. To perform inter-calibration in the AQUACOSM community,

QA measures in each of the mesocosm facility should be determined. The common QA measures that are

determined based on this SOP and the valid sampling and analysis methods used during the experiments that

shall be taken by each mesocosm facility are listed as follows:

● “Field sampling and sample labelling

● Sample storage and preservation;

● Laboratory analysis” [45].

Further suggestions provided by [45] (not specific to phytoplankton analysis) are as follows:

● “Establishing routine internal quality control

● Participation in external QA schemes” [45].

Other QA/QC practices that must be carried out

● Visual check for clumping

● Consider the settling period (before the experiment (sampling) OR during counting and

identification)

● Repeat the application of Iodine every 6 months

● Pay attention to the confidence level of microscopic enumeration of colonial or filamentous

phytoplankton species

● Consider the resolution/degree of identification and take pictures of unidentified species or species

which have unsure identification.



Table 1-5: Reagents


Ethanol (normally 70-99 %) [1] Ethanol, water

Instead of pure ethanol, cheaper methylated

spirit can be used. If mesozooplankton is

stored for molecular analysis, a solution from

pure ethanol should be used (70%).

Solvent cabinet

Formaldehyde (37 % by volume) [1] See Appendix I: Phytoplankton for detailed

information about preparing formaldehyde

solution.

Fume Cabinet/Hood

Buffered sucrose formaldehyde (Buffered

Formalin) [5]

See Appendix I: Phytoplankton for detailed

information about preparing buffered

formalin solution.

Acidified Lugol’s Iodine See Appendix I: Phytoplankton for detailed

information about neutral, acidic and alkaline

Lugol’s solution.

Fume Cabinet/Hood



1.9 References 1-Phytoplankton

[1] European Commission, “Directive 2000/60/EC of the

European Parliament and of the Council of 23

October 2000 establishing a framework for

Community action in the field of water policy,” 2000.

[2] G. WOLFRAM, K. DONABAUM, and M. T. DOKULIL,

“Guidance on the Monitoring of the Biological

Quality Elements Part B2 - Phytoplankton,” Vienna,

2015.

[3] ASTM International, “Standard Terminology Relating

to Water,” 2013.

[4] WHO, Handbook: Good Laboratory Practice (GLP):

Quality practices for regulated non-clinical research

and development, 2nd ed. Switzerland: TDR/WHO,

2009.

[5] OECD, OECD series on principles of good laboratory

practice and compliance monitoring. Paris, 1998.

[6] Council Directive, “89/391/EEC of 12 June 1989 on

the introduction of measures to encourage

improvements in the safety and health of workers at

work,” Off. J. Eur. Communities , vol. 183, no. 29, p.

8, 1989.

[7] OSHA, “European directives on safety and health at

work - Safety and health at work.” [Online].

Available: https://osha.europa.eu/en/safety-and-

health-legislation/european-directives. [Accessed:

16-Jun-2017].

[8] D. on E. and L. S. Committee on Prudent Practices in

the Laboratory, Board on Chemical Sciences and

Technology, Prudent practices in the laboratory :

handling and management of chemical hazards.

National Academy of Sciences, 2011.

[9] ECHA, “Formaldehyde - Substance Information.”

[Online]. Available:

https://echa.europa.eu/substance-information/-

/substanceinfo/100.000.002. [Accessed: 16-Jun-

2017].

[10] Ministerium Für Ein Lebenswertes Österreich,

“LEITFADEN ZUR ERHEBUNG DER BIOLOGISCHEN

QUALITÄTSELEMENTE TEIL C - ARBEITSSICHERHEIT,”

Wien, 2016.

[11] European Commission, “EU Waste Legislation.”


http://ec.europa.eu/environment/waste/legislation

/a.htm. [Accessed: 16-Jun-2017].

[12] European Commision, “COMMISSION DECISION of 3

May 2000 replacing Decision 94/3/EC establishing a

list of wastes pursuant to Article 1(a) of Council

Directive 75/442/EEC on waste and Council Decision

94/904/EC establishing a list of hazardous waste

pursuant to Article 1(4) of Council Directive

91/689/EEC on hazardous waste (notified under

document number C(2000) 1147) (Text with EEA

relevance) (2000/532/EC),” Off. J. Eur. Communities,

vol. 69, 2000.

[13] U. Mischke et al., “EU FP7 226273, WISER deliverable

D3.1-4: guidance document on sampling, analysis

and counting standards for phytoplankton in lakes,”

pp. 1–51, 2012.

[14] EPA (IRELAND), “Water Framework Directive:

Sampling Manual for Rivers and Lakes,” 2016.

[15] H. Utermohl, “Zur vervollkommnung der

quantitativen phytoplankton methodik,” 1958.

[16] UNESCO/IOC, MICROSCOPIC AND MOLECULAR

METHODS FOR QUANTITATIVE PHYTOPLANKTON

ANALYSIS. Paris: O. (IOC Manuals and Guides, no.

55.) (IOC/2010/MG/55), 2010.

[17] Aarhus Universitet – National Environmental

Research Institute (AU) and Middle East Technical

University (METU), “EU FP7 244121, REFRESH

Deliverable 3.4: Protocols for field experiments and

monitoring along a climate gradient in task 2,”

2014.

[18] P. Pitta, A. Giannakourou, and U. Christaki,

“Planktonic ciliates in the oligotrophic

Mediterranean Sea: longitudinal trends of standing

stocks, distributions and analysis of food vacuole

contents,” Aquat. Microb. Ecol., vol. 24, pp. 297–311,

2001.

[19] Baltic Marine Environment Protection Comission,

“Monitoring of phytoplankton species composition,

abundance and biomass,” 2017.

[20] IGB, “Taxaliste Phytoplankton O 2.17 | Drupal.”



[Online]. Available: http://www.igb-

berlin.de/en/project/taxaliste-phytoplankton-o-

217. [Accessed: 15-Jun-2017].

[21] Baltic Marine Environment Protection Comission,

“HELCOM Phytoplankton Expert Group (HELCOM

PEG),” HELCOM, 2016. [Online]. Available:

http://www.helcom.fi/helcom-at-

work/projects/phytoplankton. [Accessed: 05-Sep-

2017].

[22] AlgaeBase, “AlgaeBase,” 2017. [Online]. Available:

http://www.algaebase.org/. [Accessed: 19-Jun-

2017].

[23] R. A. Horner, A Taxonomic Guide to some common

marine phytoplankton. Bristol, UK: Biopress Limited,

2002.

[24] C. R. Tomas, Identifying marine phytoplankton. San

Diego: Academic press, 1997.

[25] J. Throndsen, G. R. Hasle, and K. Tangen,

Phytoplankton of Norwegian coastal waters. Oslo:

Almater Forlag AS, 2007.

[26] K. J. Flynn et al., “Misuse of the phytoplankton-

zooplankton dichotomy: The need to assign

organisms as mixotrophs within plankton functional

types,” J. Plankton Res., vol. 35, no. 1, pp. 3–11,

2013.

[27] C. A. JEROME, D. J. S. MONTAGNES, and F. J. R.

TAYLOR, “The Effect of the Quantitative Protargol

Stain and Lugol’s and Bouin’s Fixatives On Cell Size:

A More Accurate Estimate of Ciliate Species

Biomass,” J. Eukaryot. Microbiol., vol. 40, no. 3, pp.

254–259, May 1993.

[28] U. Mischke et al., “Dataset: ‘Phytoplankton’

(according to the WISER project)

www.freshwaterecology.info - the taxa and

autecology database for freshwater organisms,”

www.freshwaterecology.info. [Online]. Available:

http://linkinghub.elsevier.com/retrieve/pii/S14701

60X15000758.

[29] C. T. Kremer, J. P. Gillette, L. G. Rudstam, P. Brettum,

and R. Ptacnik, “A compendium of cell and natural

unit biovolumes for >1200 freshwater

phytoplankton species,” Ecology, vol. 95, no. 10, p.

2984, Oct. 2014.

[30] I. Olenina et al., “Biovolumes and size-classes of

phytoplankton in the Baltic Sea,” HELCOM Balt.Sea

Environ. Proc., vol. 106, no. 106, p. 144 pp, 2006.

[31] H. Hillebrand, C.-D. D?rselen, D. Kirschtel, U.

Pollingher, and T. Zohary, “BIOVOLUME

CALCULATION FOR PELAGIC AND BENTHIC

MICROALGAE,” J. Phycol., vol. 35, no. 2, pp. 403–424,

Apr. 1999.

[32] K. Olrik, P. Blomqvist, P. Brettum, G. Cronberg, and

P. Eloranta, Methods for quantitative assessment of

phytoplankton in freshwaters. P. 1: Sampling,

processing, and application in freshwater

environmental monitoring programmes. 1998.

[33] N. J. Gotelli and R. K. Colwell, “Quantifying

biodiversity: procedures and pitfalls in the

measurement and comparison of species richness,”

Ecol. Lett., vol. 4, no. 4, pp. 379–391, 2001.

[34] B. Brierley, L. Carvalho, S. Davies, and J. Krokowski,

“Guidance on the quantitative analysis of

phytoplankton in Freshwater Samples,” Edinburgh,

2007.

[35] E. Rott, N. Salmaso, and E. Hoehn, “Quality control of

Uterm?hl-based phytoplankton counting and

biovolume estimates?an easy task or a Gordian

knot?,” Hydrobiologia, vol. 578, no. 1, pp. 141–146,

Mar. 2007.

[36] E. Hoehn, H. A. M. Ketelaars, and B. Ewig, Erfassung

und Bewertung von Planktonorganismen:

Arbeitsgemeinschaft Trinkwassertalsperren eV

Arbeitskreis Biologie. Oldenbourg, 1998.

[37] J. Padisák and R. Adrian, “Biovolumen,” in

Biologische Gewasseruntersuchung, Band 2., Gustav

Fischer Verlag, 1999, pp. 334–368.

[38] D. Hepperle, “OPTICOUNT - Software for the

enumeration of plankton,” 2005.

[39] A. Lyakh, “The free form deformation of phyto-

plankton models,” in Lecture Notes in Computer

Sciences 2331, 2002, pp. 192–201.

[40] K. Vuorio, L. Lepistö, and A.-L. Holopainen,

“Intercalibrations of freshwater phytoplankton

analyses.,” Boreal Environ. Res., vol. 12, no. 5, 2007.

[41] SMHI, “Nordic Microalgae.” [Online]. Available:

http://nordicmicroalgae.org/. [Accessed: 19-Jun-

2017].



[42] HELCOM, “HELCOM Phytoplankton Expert Group

(HELCOM PEG).” [Online]. Available:

http://helcom.fi/helcom-at-

work/projects/phytoplankton/. [Accessed: 19-Jun-

2017].

[43] Alfred Wegener Institute for Polar and Marine

Research, “PLANKTON*NET.” [Online]. Available:

http://planktonnet.awi.de/. [Accessed: 19-Jun-

2017].

[44] CEN EN 15204, “Water quality–Guidance standard

for the routine analysis of phytoplankton abundance

and composition using inverted microscopy

(Utermohl technique),” 2006.

[45] European Commission, “Common Implementation

Strategy for the Water Framework Directive

(2000/60/EC) Monitoring under the Water

Framework Directive,” Luxemburg, 2003.

[46] İ. Tuney and M. Maroulakis, “PHYTOPLANKTON

SAMPLING METHODS,” 2014.

[47] P. Santhanam, A. Begum and P. Pachiappan (ed.).

Basic and Applied Phytoplankton Biology. Springer

Singapore, 2019.

Co-funded by the European Union D4.1 Standard Operating Procedures| 2 Zooplankton


2. Zooplankton


Biomass the amount of living matter present in the mesozooplankton sample [3]

Epilimnion or

Upper mixed layer

water above the pycnocline, i.e. thermocline (freshwater and marine systems)

or halocline (marine) a in a stratified body of water [1]

Fixation protection from disintegration of the morphological structure of organisms [1]

Halocline vertical zone in the oceanic water column in which salinity changes rapidly with

depth [4]

Hypolimnion or

deeper layers

water below the pycnocline, i.e. thermocline (limn./mar) or halocline (mar) a in

a stratified body of water [1]

Littoral zone shallow marginal zone of a body of water within which light penetrates to the

bottom; usually colonised by rooted vegetation [1]

Pelagic zone body of water beyond the littoral zone [1]

Plankton organisms drifting or suspended in water, consisting chiefly of minute plants or

animals, but including larger forms having only weak powers of locomotion [1]

Sampling site

(Sampling station)

general area within a body of water from which samples are taken [1]

Stratified water freshwater (generally lakes/standing water) or marine waters with a strong

density gradient (normally temperature and/or in marine systems salinity)

resulting in an upper, normally warmer, mixed/isothermal layer floating on a

denser, usually colder and or more saline, also isothermal water

Subsampling collection of a sub-sample that consists of a known fraction of the total sample

and that is representative of the quantity and species composition of the latter

[1]



Thermocline

(metalimnion)

layer in a thermally stratified body of water in which the temperature gradient

is at a maximum [1]

Zooplankton animals (heterotrophic organisms) present in plankton [1]


Different preserving solutions with different areas of application are available in the literature. Some of the

preserving reagents are summarized in Table 2-1.

Table 2-1: The materials and reagents used in analysis of mesozooplankton


Ethanol (normally 70-99 %) [1] Ethanol, water

Instead of pure ethanol, cheaper methylated spirit can be used. If mesozooplankton is stored for molecular analysis, a solution from pure ethanol should be used (70%).

Solvent cabinet

Formaldehyde (37 % by volume) [1] See Appendix II-A for detailed information

about preparing formaldehyde solution.

Fume Cabinet/Hood

Buffered sucrose formaldehyde (Buffered Formalin) [5]

See Appendix II-A for detailed information

about preparing buffered formalin solution.

Acidified Lugol’s Iodine See Appendix II-A for detailed information

about neutral, acidic and alkaline Lugol’s solution.

Fume Cabinet/Hood

In addition, different narcotizing agents such as Tricaine Methanesulfonate (MS-222), magnesium chloride,

carbonated water, chloroform and methyl alcohol are commonly used in marine sampling [6].

According to [1], preserving solutions should meet the following requirements:

● The most frequently used preservatives in mesozooplankton research used to be Lugol’s Iodine in

freshwater and formaldehyde in both freshwater and marine systems, while ethanol is presently



increasingly used for fixation, because it presents less health hazards and allows genetic analysis.

One should note that formaldehyde can only be used with ‘special permission’ in some laboratories

as it is associated with allergies and cancers. For that reason, it should be handled with care and

authorized personal should be contacted for permission and guidance before use.

● The effect of the fixative (concentration, method of addition) on the discoloration, deformation and

even loss of organisms by chemical shock or otherwise, should be considered before choosing

fixative. Alternatively, with increasing use of live analysis of zooplankton – this can be evaluated to

avoid fixation altogether. This would also allow other chemical analyses of the same organisms down-

stream. This will be further discussed in the next version of this SOP.

● The preservative should effectively prevent the microbial degradation of organic matter for at least

the storage period of the samples.

● The preservative should guarantee a good recognition of taxa for at least the storage period of the

samples.

● The preserving solutions should be kept in closed bottles, which should be stored in a fluid tight

plastic box or container during transportation to and from the sampling (to allow fixation directly

after sampling and to prevent spillage of fixative. Use of gloves and pipettes to transfer the solution

to the plankton samples are recommended [1]. Most fixatives, such as Lugol’s Iodine and formaline

need to be renewed after certain time in storage, typically due to oxidation. Oxidation rate depends

on fixative, buffer and storing conditions. Samples should be stored dark and cold (but above

freezing) for best preservation.

● The advantages and disadvantages of each of these solutions are detailed in Appendix II-A. It should

be noted that the reagent may affect the size of the organisms (shrinkage). The shrinkage may differ

depending on fixative concentration and status of the fixed organisms before fixation. Please see

Appendix II-A for more information.

●



In this section, general guidance on the protection of health and safety while sampling, analysing and

counting mesozooplankton from mesocosm experiments will be provided to minimize the risk of health

impacts, injuries and maximize safety. The users of this SOP are expected to be familiar with the Good

Laboratory Practice (GLP) of World Health Organization (WHO) [7] and Principles on GLP of Organisation for

Economic Co-operation and Development (OECD) [8]. Health and Safety Instructions of the mesocosm

facility, if there are any, shall be followed properly to protect the people from hazardous substances and the

harmful effects of them.


site or in the laboratory), the work should be practiced consistent with national and EU regulations (see the







OSH Framework Directive 89/391/EEC, [9]). Other regulations and guidelines can be found in EU – OSHA


shall be taken by the users of this SOP and the scientist-in-charge shall ensure that those measures comply


The table below summarizes the hazards and risks and the measures for preventing them associated to the

laboratory studies carried out on mesozooplankton:


Occupations at

risk


Laboratories o Risk of inhaling

chemicals that are

used for

mesozooplankton

sample fixing

o Accidental spills

o Use of fume cabinets/hood


o Appropriate personal protection (protective

gloves) and hygiene measures

o Storage of samples in glass bottles to minimize

diffusion of the fixative


o Use of closable cooling boxes to transport

samples between field and laboratory

o Using of closable plastic boxes to store the

sample bottles

2.3.2 Safety Instructions for Sampling

Please see the Water Chemistry SOP (Deliverable 4.1.4) for safety instructions for sampling.

As a best practice advice, if the samples are collected from a boat, always have a shore-based contact in case

of emergencies. Check the weather forecast to ensure safe and effective working conditions. For safety

reasons, it is recommended that field work should not be undertaken by unaccompanied persons, but by a

minimum of two people [1].

2.3.3 Working and Personal Protection

Please see the Water Chemistry SOP (Deliverable 4.1.4) working and personal protection equipment

suggested for use in water sampling.





2.3.4 Use, storage and disposal of reagents and chemicals

The user of this SOP needs to visit the Safety Data Sheets (SDSs), which is provided by the manufacturer for

any chemicals. The SDSs contain necessary information for protection before using, storing and disposing the

reagents as well as other chemicals. Only experienced personnel should be responsible for the use of

reagents, preservatives and chemicals in a way compatible with the laboratory rules.

Important: Use of formaldehyde as a fixative can result in “the generation of bis(chloromethyl)ether” - that

is a potential human carcinogen [11]. In addition, formaldehyde, is an irritant and can enter the body via

inhalation and damage tissues seriously. For more information on formaldehyde, see the substance

information [12] provided by European Chemicals Agency (ECHA).

2.3.5 Use, storage and disposal of equipment

Zooplankton sampling equipment needs to be cleaned in between sampling events [13]. The storage of

equipment should be compatible with the operating manual/instructions for use. Cleaning, disinfection and

storage of equipment should be taken care under the risk of spreading parasites, diseases, foreign species,

and pathogens.

2.4 Environmental Indications

A plan for the disposal of mesocosm waste needs to be prepared prior to the experiments. The plan must

comply with the EU Waste Legislation [14] and The List of Hazardous Wastes [15] provided by the European

Commission. The Safety Data Sheets (SDSs) need to be followed for the disposal of reagents and chemicals

prior to waste disposal.

2.5 Sampling Zooplankton

2.5.1 Prior to sample collection

The AQUACOSM Standard Operating Procedure (SOP) on sampling, analysis and counting of

mesozooplankton needs to be reviewed by the research team prior to initiating the sampling. The sampling

method to be employed, sample type and volume, equipment and supplies needed for sample collection

must be identified prior to the start of a sampling event based on the objectives and main questions of the

study. In addition, the type of mesozooplankton organisms such as larger crustaceans or smaller ones like

rotifers, and larval meroplankton may require different methods for sampling, counting and analysis [2].

2.5.2 Required Strategies

As suggested in [1] and [5], the documentation that should be available before the start of field work is listed

below:







● Sampling plan; including the description of objectives and strategy and methods, data QA/QC

objectives – (See SOP on QA/QC, Deliverable 4.1.6)

● Applicable safety instructions

● The registration (metadata) forms (provided in Appendix II-B).

2.5.3 Preparation and calibration of sampling equipment

The sampling equipment should be checked and prepared in time before the sampling and left in proper

order for the next sampling. The required actions to prepare the equipment for mesozooplankton sampling

are summarized from [1] as follows:

2.5.3.1 Overview of equipment and instruments

● Nets, especially finer meshed (< 200 µm) should be washed “in warm freshwater with detergent or

in an ultrasonic water-bath”, to minimize the risk of clogging and to ensure optimum filtration

capacity [1], especially after waters with high concentrations of large diatoms or other organisms

that can stick to the mesh and permanently clog the net if dried.

● Before use, plankton nets and the draining cups should be visually checked for tears or holes.

● The string (rope) should be checked to ensure that it is securely attached to the sampling equipment

(plankton net/volume sampler). It should be visually checked for the depth marking on the line or

the winch.

● The closing mechanism of the sampler should be checked to ensure that it is well functioning.

● To prevent cross-contamination of organisms between mesocosms, the sampling equipment should

be cleaned thoroughly between uses in the different waters.

2.5.3.2 Sample bottle preparation and preservation

In order to ensure minimal change of the sample content due to e.g. predation of smaller organisms in the

concentrated sample (codend feeding) it is suggested to fix the samples as soon as possible after sampling.

This can either be done by adding fixative to the sample bottles before commencing the fieldwork. If it is not

possible to add fixatives to the bottles (e.g. due to restrictions of use of toxic chemicals at the site), an

alternative is to add non-toxic narcotizers to the bottle to prevent condend feeding before fixing. It is also

suggested to mark the bottles before sampling (see Appendix II: Zooplankton for details on labelling). To

ensure safe marking, it is suggested to add a water-resistant paper with sample identification written with a

pencil inside the bottles.

For more information on the effectiveness of the devices for specific type of mesozooplankton, one can visit

Welch (1948) [16], Edmondson and Winberg (1971) [17], de Bernardi (1984) [18], DeVries and Stein (1991)

[19], Harris et al. (2000) [20], Schwoerbel (2016) [21], Manickam et. al. (2019) [22] and Santhanam et. al.

(2019) [23].



2.5.4 Sampling Equipment

Figure 2-1: Plankton nets Different types of plankton nets: a) definition of net length and opening diameter and demonstrating how the filtering surface is increased by a cylindrical net section above the conical part, b) conical plankton net, c) cylindrical plankton net (Indian Ocean Standard Net), and d) a net with Apstein cone (Adopted from [1] and [24]).

Zooplankton sampling from mesocosms can be performed for qualitative and quantitative analysis.

Qualitative mesozooplankton sampling mainly provides information about “the species composition, number

of species, size distribution and relative dominance of species and groups of mesozooplankton” [1]. On the

other hand, quantitative sampling provides information on the quantity of mesozooplankton per unit volume

of water, as well as a basis to calculate the total biomass for the mesozooplankton and individual

mesozooplankton species [1]. In addition, size diversity can be estimated based on quantitative analysis

carried out. Whenever there are no significant differences in effort of sampling, it is of the latter reasons

recommended to conduct fully quantitative sampling.

There are many different devices to sample mesozooplankton; including bottles, traps, tubes, pumps and

nets. The effectiveness of these different techniques varies according to the type of mesozooplankton [2],

type of the mesocosm (depth of strata), type of the samples (point or integrated), volume of water to be

sampled and the studied problem [18].

2.5.4.1 Plankton Nets

Plankton nets, have been the most common tool to sample zooplankton for soon 200 years. They are

available in various dimensions and mesh sizes that vary according to the study objectives (e.g. Fehler!

Verweisquelle konnte nicht gefunden werden.). However, drawbacks with plankton nets is that they suffer

from variable rate of under-sampling due to the resistance for water to pass through the meshes, and thus

“loss of organisms through the meshes, net avoidance, and variation in filtration efficiency” [6]. The degree

of under-sampling generally increases with decreasing mesh size and increasing amounts of objects caught

by the net. “Net efficiency is affected by a series of factors including characteristics of the fabric used to

a b c d



construct the net (i.e. the gauze), mesh sizes, porosity, speed of sampling, avoidance by target organisms,

escape of sampled organisms, and clogging” [18]. Moreover, net sampling only measure the average density

of organisms integrated over a certain volume but cannot measure the density or distribution of organisms

on small spatial scales [6].

In order to optimize the efficiency of water filtering, plankton nets should have a large filtering surface

relative to their opening. This can be achieved by a long net compared to the opening (i.e., opening diameter

of 30 cm and length of about one meter), as recommended by [1]. A larger filter surface can also be achieved

with a shorter net by using a cylindrical net section above the conical part in comparison with a conical

plankton net with the same opening diameter and length [1] (compare Fehler! Verweisquelle konnte nicht

gefunden werden.a-c), and may also be achieved by decreasing the opening compared to the net surface

area as with an Apstein-cone (Fehler! Verweisquelle konnte nicht gefunden werden.)

According to the size classes of the mesozooplankton, appropriate mesh sizes can be selected.

The appropriate mesh size for the plankton nets to be used qualitative sampling are summarized in Table 2-3

below.

For higher sampling efficiency, (i.e. a balance between catching small sized organisms with limited escape

ability, and faster swimming larger sized plankton), use of different nets with different mesh sizes and hauling

speed, is recommended. NOTE that the size of most mesocosms do not allow substantial sampling of larger

meso- and macrozooplankton as their abundances are too low to allow removal in numbers adequate for

statistical analyses without exhausting the content of the mesocosms. In such cases it is recommended to

sample these by emptying the mesocosms through a net at the end of the experiment.

Table 2-3: Appropriate mesh size for qualitative sampling of different sizes of mesozooplankton

Targeted plankton Size (µm) Mesh Sizea Notes on the nets with specified

mesh sizes

Microzoo-plankton

20 - 200 Typically 20-90 µm, in

Freshwater 90 µm

nets with Apstein

collars are commonly

used for freshwater

crustaceans, and 40-

50 µm to include

Rotifers.

+ Useful in oligotrophic waters

- Are expected to have clogging

problems, that can be partially

ameliorated with an Apstein-collar.

90 µm is a common compromise

between filtering efficiency and

small mesh size.

Mesozooplankton 200-2000 200 μm (or 150 - 180

µm) mesh is used to

+ Useful in sampling rotifers and

crustaceans larger than 200 µm, but



catch all

mesozooplankton,

including predatory

species in freshwater

many rotifers and early crustacean

development stages are technically

microzooplankton (<200 µm) and

will be strongly under-sampled.

- Low efficiency in sampling both

types 500-2000 300 - 350 µm

Macrozooplankton >2000 1000 µm or larger,

with non-filtering

codend for delicate

macrozoopankton,

such as medusae.

Slow swimming macrozoopankton

such as medusae can be collected

with e.g. 1000 µm mesh nets, often

with large non-filtering codends to

minimize damage. However, fast

swimming “zooplankton” like krill is

caught with larger and faster devises

like Isaak-Kidd trawls. But such

animals are not generally possible to

hold in mesocosms.

a Mesh sizes mentioned should be regarded as for guidance. Mesh sizes will vary somewhat from manufacturer to manufacturer.

2.5.4.2 Sampling with plankton nets:

Nets and the buckets to be used for sampling should be cleaned, checked for holes and tears prior to

sampling. Then, the collection bucket is attached to the cod end of the net. The bridled end of the net should

be attached to a string (rope) with markings for determined depths, to measure the sampling depth.

The net should be retrieved by pulling it back to the surface with a steady constant speed, ca 0.5 m/s or

slower if clogging is severe for smaller meshed nets. When at the surface, the catch can be rinsed into the

codend by dipping the net and rapidly lifting it out of the water (keeping the opening above water at all

times).

When depth stratified samples are needed nets with closing mechanisms may be used, such as Apstein

closing nets [24], or a Clarke-Bumpus sampler [25] that is a plankton net connected to a flowmeter, allowing

measurement of the volume of water filtered by the sampler. For additional information and best practice

advices on using Clarke-Bumpus plankton samplers, please refer to [18] and [26].



Then, the contents of the plankton net should be rinsed to the collection bucket by spraying water against

the outside of the net. The collection bucket should be removed from the net vertically. The contents in the

collection bucket can be concentrated by swirling the bucket.

Risk of contamination through net sampling: Plankton nets represent large surfaces and are therefore prone

to cause contamination among mesocosms and samples. Proper rinsing after every single sample is

mandatory in order not to contaminate subsequent samples and mesocosms (i.e. animals that are not

washed from the net after a net haul may end up in the mesocosm sampled/in the sample form the

mesocosm sampled next). Contamination is particularly problematic if mesocosms contain different

communities, which is an issue in experiments involving manipulation of connectivity. Note that protists and

bacteria will easily be transferred among mesocosms through net sampling. To prevent cross-contamination

of organisms between mesocosms, the sampling equipment should be cleaned thoroughly between uses in

the different waters. Moreover, in some mesocosms, there are sampling ports to avoid contamination among

mesocosms.

Using plankton nets for quantitative sampling, one shall measure and keep record of the quantity of water,

that passes through the net. The volume of filtered water should be calculated with the following

formulation, as given in [18]:

𝑉 = 𝜋 × 𝑟2 × 𝑑

where V is the volume of water filtered by the plankton net, r is the radius of the mouth of the net, and d is

the distance through which the plankton net is towed.

De Bernardi (1984) [18] underlined the errors in the given calculation that can occur due to clogging of the

net. Accordingly, use of nets with flow meters is strongly recommended, both for freshwater and estuarine

mesocosms [6]. The best position for the flow meter was suggested to be in a position midway between the

center and the net rim [27]. Moreover, a second flow meter outside the net can provide an estimate of net

speed, and the two meters combined can yield an indication of filtration efficiency and clogging.

In case of using a flow meter, the volume of water that is sampled can be calculated using the specific

formula, generally provided by the manufacturer of the flow meter [6].

2.5.4.3 Using water samplers to collect zooplankton in mesocosms

Several types of water or volume samplers (water bottles, tube samplers and pumps) can be used to collect

(nano-micro- and) mesozooplankton while removing the zooplankton together with a known volume of

water that will later be filtered to separate the mesozooplankton from the water sample. This approach has

several advantages compared to net samples: 1) it does not change the relative composition of the

organisms in the ecosystem by selective removal of organisms in the mesocosm, 2) the sampled volume can

be precisely determined, and 3) for two of the water sampling types discussed below (water bottles and

pumps) samples can be taken at higher spatial resolution than with most nets.

Water bottles (Figure 2-2) sample relatively small amounts of water. Water bottles are thus generally

“efficient for small, less motile organisms” which can “yield low collection efficiency when used to sample



the larger, scarcer, and more active zooplankters” [18]. Thus, as relatively small amount of water is sampled,

the number of replicates generally needs to be higher, especially to account for spatial heterogeneity of

mesozooplankton distribution horizontally and vertically [18]. However, an alternative way to minimize

spatial heterogeneity in mesocosms, is to mix the mesocosms thoroughly before sampling, or pooling

samples from several locations in the mesocosms. Note that analysis time and cost can be reduced if single

samples are mixed [1], rather than counted separately.

Figure 2-2: Examples of volume samplers that can be used for water and (zoo)plankton sampling: a) Ruttner bottle, b) Friedinger bottle [1], c) Niskin water samplers, and d) Van Dorn samplers [28], [29]. All have closure mechanisms that allows sampling at desired depths [6].

Tube samplers can be seen as a “long water bottle” used to collect an integrated sample when lowered

through the water column [18]. Using (rigid) tube samplers, the entire water column can be sampled (i.e.

from the surface to e.g. a few cm above the sediment) with an appropriate diameter in shallow water

mesocosms ([30], [31]). They include models such as Heart Valve and Limnos Samplers, described in Knoechel

and colleagues (1992) and Pennak (1962) ([32], [33]). When using rigid tube samplers, one should consider

that the length of the tube allows for sampling the wanted depth without being to long for easy handling

[18]. It is also recommended in [1] to choose a sampler that is transparent (e.g. Plexiglas) to have a better

idea for what and how much water is being sampled, allows the free flow of water when lowered and has a

rapid and tight closing system.

When sampling, the tube should be lowered vertically to the predetermined depth at an even and moderate

speed to eliminate most of the pressure wave in front of the opening [1]. The tube is then retrieved at a



moderate speed. If the volume samplers are not used with an integrated net (ex. Bottles), the water should

be filtered through a plankton net (or other straining equipment).

To overcome heterogeneity in the mesocosms, it is advisable to pool samples from several locations in the

mesocosm. The (sub)sample(s) should be filtered through a net with an appropriate mesh size, making sure

that all material on the inside of the net is rinsed into the draining cup. The material should be transferred to

the sample bottles and preserved or analysed as soon as possible after collecting on the net.

Excess water could be returned to the mesocosm if volumes are limiting, but returning water to the

mesocosm that is sampled by nets may also introduce bias in organismal composition, that should also be

considered.

Plankton pumps (hand pumps and motorized pumps), described e.g. by Nayar et al (2002) and Waite &

O’Grady (1980) ([34], [35]) offers an alternative method to sample either at specific depths or over depth

intervals in mesocosms. Sampling procedures and how to effectively use pumps to collect mesozooplankton

were also summarized in De Bernardi (1984) [18], as quoted from Tonolli (1971) [26]. Compared to other

volumetric samplers, pumps “permit the collection of the largest volumes of water” [18]. Accordingly,

pumps can be employed as an alternative to bottles and tubes with some advantage if large water volumes

are needed. However, “large active plankton species are liable to be sampled less efficiently using a

plankton pump than by other types of quantitative samplers. The opposite can be the case for small species”

[1], [18]. Use of “a large plastic funnel (diameter about 50 cm) at the end of the sampling tube” is

recommended “to prevent escape of jumping copepods” [1]. Motorized plankton pumps with continuous

flow-through are recommended rather than hand-powered plankton pumps, because motorized pumps

provide a regular flow, thus providing better estimates of the quantity and composition of the plankton [1].

The approach used for quantitative estimation and small-scale distribution of plankton with pumps have

been summarized in Kaisary and colleagues (2012) as: “an open-ended inlet hosepipe is lowered into the

water and the outlet pipe is connected to a net of suitable mesh size. The net is particularly submerged in a

tank of a known volume. This prevents damage to the organisms. The mesozooplankton is filtered through

the net. A meter scale on the pump records the volume of water filtered” [6].

Sampling by gravity, can be used to collect mesozooplankton. Some land based mesocosm facilities allow

taking water samples from a port or by a hose through gravity. In this case a sufficient amount of water needs

to be collected (depending on density of organisms). The water sample is screened by an appropriate mesh

(50-100µm) in the lab. Given that mesozooplankton is not retained quantitatively in the mesh, the screened

water can be used for other analyses, e.g. chlorophyll-a and nutrient analyses, or may be added back to the

mesocosm, if needed.

2.5.4.4 Additional materials generally needed for zooplankton sampling

Weight (for net sampling only, non-toxic materials should be considered in mesocosms). Squirt/Wash bottle

to rinse out the net and the draining cup. Small plastic funnel for transferring the water sampled to sample

bottles. Plastic (bucket) for pooling multiple water samples (e.g. from volume sampler). Filtration equipment



to collect or concentrate samples from volume samplers. The filtration equipment may be either a plankton

net, with an appropriate size to filter the targeted organisms or “a large funnel with draining cup fitted with

a netting” [1]. Gloves (rubber or vinyl), waterproof notebook, waterproof pen, if ethanol or other solvents

is used, an alcohol-proof pen or pencil is recommended for both internal and external marking, pipettes (to

add preservative)

2.5.4.5 Sampling Containers and Sample Bottles

Bottles (100 ml, 200 ml or 250 ml), recommended to be (brown) bottles with screw-tops or glass vials for

storing samples [1]. Select the right type of bottle for samples based on the type of the reagent used. For

instance, plastics are not suitable for storing samples if Lugol’s Iodine is used as preserving solution [1]. See

Appendix II-A for more details.

Labels, recommended to be from waterproof paper. As a best practice advice, mesocosm name,

(treatments) date and time of sampling, the sampling depth, the sampling equipment used should be stated

on the labels of each mesozooplankton sample [2]. Volume of water filtered, and the mesh size of the net

used in filtering should also be recorded on the labels.

2.5.5 Sampling Design for Shallow and Deep Mesocosms

As a best practice advice, if the study aims to estimate species composition and abundance in the mesocosm,

samples should be taken both from different depths of whole mesocosms; as the vertical and horizontal

distribution of mesozooplankton is uneven.

Moreover, “if the objectives of the study require information to be collected regarding spatial variation in

general, or a high level of accuracy in the estimates, it will be necessary to draw up a sampling design

(programme) that is adapted to the” width and the depth conditions of the mesocosm [1].

For instance; “stratified, random sampling (see [36] for more information) of mesozooplankton is based on

dividing the mesocosm horizontally and vertically into a series of sampling units (strata), from which a

random selection of units to be sampled is selected. The optimum number of samples per stratum will

depend upon three factors: the size of the stratum, the variability within the stratum and the cost of sampling

in the stratum [1] ”.

In general, sampling should be carried out at an adequate distance from mesocosm walls and one shall be

careful not to disturb the plants, if available.



2.5.5.1 Sampling time and frequency

Sampling time and frequency should be determined according to the main purpose of the study. There are

many recommendations about time and frequency, such as

• According to EN 15110 (2006), mesozooplankton should normally be sampled between 10 a.m. and

4 p.m. For studies of vertical migration, sampling every 6 hours is recommended, but as a minimum,

sampling should be performed at midday and midnight, or at least roughly the same time each time

to maintain consistency and allow comparison [1] .

• “Many common species of macromesozooplankton (such as Chaoborus, Mysis) may only occur in the

water column at night. If collection of these species is desirable, it is essential that sampling occur at

least one hour after sunset” but it is best at midnight [2].

2.5.5.2 Shallow mesocosms

If mesocosms are mixed continuously, one composite (pooled) water sample (i.e., samples from different

locations and depths can be combined) can be considered as representative for the entire mesocosm. If it is

unclear whether the whole water volume is efficiently mixed, make sure to include all (thermally or salinity-

based) stratified zones be covered.

For mesocosms containing sediment and possibly with macrophytes, a detailed sampling design for

collecting samples at multiple horizontal and vertical positions might be needed, as “large-bodied

mesozooplankton (typically cladocerans) often aggregate within submerged plant stands or sediment during

daytime (Timms & Moss (1984) [37]; Lauridsen & Lodge, 1996 [38], Tavşanoğlu et al. (2012), [39]), as

macrophytes might serve as a refuge for mesozooplankton. In such mesocosms, mesozooplankton samples

should be taken from both areas containing aquatic vegetation and from areas with little vegetation, just

above the sediment surface without disturbing [1].

A suggested approach is summarized in [31] as follows:

“The entire water column, from the surface to approximately 5 cm above the sediment, is sampled from

three positions in each enclosure: 10, 30 and 60 cm from the enclosure wall. Two samples are prepared: one

to be used for chemical and phytoplankton analyses where water is sampled without touching the plants –

and one to be used for mesozooplankton analyses, where water is sampled also close to the plants”.

As sampling device for shallow and macrophyte-covered mesocosms, use of a volume sampler, plankton

pump or flexible tube (i.e. core/tube sampler) with an adequate diameter (i.e. > 6 cm) to sample from the

entire water column are recommended [1]. If tube sampler is not available, samples can be taken with

“Ruttner water sampler from the surface (i.e. 20 cm below the water surface), middle and the bottom (i.e.

20 cm above the sediment). The sample in the middle should be adjusted according to the enclosure type

and actual water depth” [31]. In addition, a heart valve or Limnos sampler can be used for sampling.



2.5.5.3 Deep mesocosms

If the water column in the mesocosm is stratified or partially mixed, the sampling procedure must be

determined after careful considerations on the water layer(s) to be sampled (e.g. sampling discrete water

layers; sampling multiple depths with subsequent pooling to one combined representative of a layer).

✓ As best practice advice, a pilot study prior to actual study can be carried out to examine

mesozooplankton densities at several horizontal and vertical sites in a mesocosm (two or three

locations previously determined, in line with the purpose of the study. The results of the pilot

study can be used in planning the actual sampling design.

✓ Sampling design and the estimation of mesozooplankton sampling precision are reviewed by

Green (1979) [40], Downing et al. (1987) [41], Eberhardt and Thomas (1991) [42], Pace et al.

(1991) [43], and Prepas (1984) [44] in more details.

✓ Absence of vertical temperature profiles indicates vertical mixing, but NOT necessarily

homogenous distribution of motile organisms like zooplankton. Heterogeneous distribution of

mesozooplankton should be assumed as the mesozooplankton densities and species

composition vary vertically and horizontally in water bodies and has not been proven to be

(practically) homogenous.

✓ Diel vertical migration (DVM) has been shown to be an important anti-predator defense

mechanism in deep stratified mesocosms (in Gliwicz (1986) [45] and Lampert (1989) [46]) that

can be taken into account for sampling.

According to the European standard EN 15110 (2006), “samples should be taken at intervals of no more than

2 m in the epilimnion and metalimnion” [1] . Accordingly, in deep mesocosms (i.e. 20 m depth), the samples

taken should represent the mesocosm vertically and horizontally. The composited samples (i.e. integrated

samples taken from a given range of depths) should include the epilimnion and the hypolimnion. An example

methodology is summarized in Berger and colleagues (2010) [47], for sampling mesozooplankton from deep

mesocosms.

As a sampling device, for deep mesocosms, it may be more appropriate to use sampling equipment that

allows efficient sampling of the whole water column, such as plankton net with a closure or a plankton pump

rather than volume samplers, especially for practicality and safety issues [1] . If vertical net hauls are used,

all depths should be covered. Remember that the net can be clogged easily in case the mesocosm includes

massive quantities of algae and other particles. However, for sampling the entire water column, a flexible

hose as tube sampler may be preferable to all alternative methods due to highest sampling efficiency (Walles

and Nejstgaard, in prep).

2.5.5.4 Best Practice Advices on selecting the right sampling instrument

In order to provide a pathway to select the right sampling equipment for quantitative mesozooplankton

sampling, De Bernardi (1984) compiled the following table from several comparison studies (Table 2-4). For



further information on the selection of the appropriate instrument, Gehringer & Aron (1968) [27], Tonolli

(1971) [26] and Bottrell et al. (1976) [48] should be visited [18].

Table 2-4: “Schematic recommendation for the choice of sampler to be used for the assessment of mesozooplankton population density under various conditions” (Redrawn from [18]).

Deep and pelagic Shallow and littoral

Point

samples

Vertically

integrated

samples

Point

samples

Vertically

integrated

samples

Samples

within

vegetation

Ruttner, etc. + - + - +

Traps ++ - ++ - +

Tubes - ++ - ++ ++

Pumps ++ + ++ - ++

Nets - ++ - - -

Plankton

samplersa - ++ - - -

a i.e. Clarke-Bumpus

2.5.6 Best practice advice on preservation and storage

Samples for quantitative analyses should be immediately preserved. If this is not possible,

1) non-preserved samples should be kept cold (2 - 5 °C) and dark as short as possible, as larger

organisms may alter the content rapidly by artificial predation rates in the concentrated sample [1].

2) preserving fluid can be added to the sampling bottles in advance. If preserving fluid has not been

added to the sampling bottles in advance, it should be added immediately after the samples have

been taken.



3) To minimize shrinkage and muscular tension, zooplankton organisms may be anaesthetized before

preservation, e.g. with carbon dioxide in mineral water.

Ethanol is a preferred fixative as it is less toxic than e.g. formalin and allows better molecular analysis of the

samples. If ethanol is used, the preserved sample should have an ethanol content of at least 70 % to 75 %.

Samples for genetic analysis based on DNA extraction should be stored in a refrigerator or freezer, but in

such a way that the sample itself does not freeze, which is ensured by preservation in high concentrations of

ethanol [1]. For such samples, ethanol with a concentration > 90 % is recommended (an ethanol content

higher than 90 % is recommended). An aqueous solution based on pure ethanol should be used for this

purpose (instead of methylated spirit).

Note that samples for genetic analysis based on electrophoresis should be stored in a freezer (approximately

-75 °C) and should not have any preserving solution added to them [1].

✓ A high ethanol concentration helps to maintain body shape (Cladocerans) as well as preventing

the animals from releasing their eggs.

If formaldehyde is used, the preserved sample should have a formalin concentration of around 4 %, i.e. 20

ml aqueous 20 % formalin should be added per 100 ml sample volume, or approximately 10% if concentrated

formaline (37%) is used.

✓ Preservation in a cold solution of formalin (6 °C) with added sucrose reduces the chances of the

animals releasing their eggs.

If Lugol’s Iodine is used, add in the amount of 0.5 ml to 1.0 ml per 100 ml sample volume. While Lugol’s has

the advantage of being non-toxic for humans, it is also a weak preservative. Thus, samples containing large

quantities of animals and other organic material (particularly samples collected in littoral regions) require

addition of more Lugol’s Iodine than samples that contain little organic material; e.g. 3 ml to 5 ml per 100 ml

sample volume. Samples preserved with Lugol’s Iodine should always be stored in the dark and preferably

chilled to below 5 °C, unless they are to be analysed within a week, in which case they can be stored in the

dark at room temperature [1]. Samples that have been preserved with Lugol’s Iodine should be straw colored

and should be checked after a couple of days for oxidation” [1]. The preservative should be renewed every 6

months for better storage.

2.6 Quantitative Analysis of Zooplankton

2.6.1 Sample Preparation

2.6.1.1 Subsampling

Zooplankton samples frequently contain many organisms and it is often impractical to count every individual.

A number of methods have been developed to subsample mesozooplankton collections.



An example subsampling process by pipettes was summarized in [49] as follows:

1. Use a swirling flask and Hensen-Stempel pipette (i.e. the pipette, which is a wide bore piston

apparatus; ensuring the organisms not to be selected against based on size, that can capture a 2.5 to

50 ml volume of liquid).

✓ Edmondson and Winberg (1971) suggest that the opening of any pipette used to take

subsamples, whether automatic or manual, should exceed 4 mm [17].

✓ Since the volume of the pipette is fixed, the volume of the sample should be altered to give

subsampling densities > 60 individuals per subsample.

2. The sample is accurately brought to the desired volume and then poured into the flask.

3. It is then swirled in a figure eight pattern until mixed.

4. While in motion, the Stempel pipette is inserted into the flask and the subsample is taken.

✓ In [48], it is found that the coefficient of variation for subsampling in this way stabilized at

0.08 when the density of organisms in the subsample exceeded 60 individuals.

Another subsampling process by Plunger sampler is as follows:

Figure 2-3: The Plunger Sampler (Adopted from [50])



“The sample is filtered at an appropriate mesh size, which is smaller than the one used in the field (for

instance, 20 µm). The content is rinsed into a sampler (as in Figure 2-3) and it is filled until 100 ml.

The piston (Plunger) is put into the sampler (the tube). Make sure that the plunger sampler is placed

vertically and straight in the sampler and the lid is tight. The water in the sampler is mixed carefully

and gently by turning the sampler upside down a number of times. Place the sampler on the table

and drag up (gently, but quickly) a known volume (i.e. 2.5-10 ml) of sample. Depending on the density

of the animals, several subsamples, up to 40 ml in total, could be taken from one sample and added

into one counting chamber. Pour the subsample in a counting chamber and rinse the piston into the

counting chamber” (Personal communication with Liselotte Sander Johansson, AU, and from [50]).

A wide variety of plankton splitters have also been developed for the subsampling of mesozooplankton

samples (e.g. Kott (1953) [51]), which are generally made of plastic with internal partitions. Folsom plankton

splitters are widely used (in marine research) and by which, the mesozooplankton samples are divided into

equal fractions. Kaisary and others (2012) summarized the subsampling methodology with a plankton splitter

as follows:

“The mesozooplankton sample to be sub-sampled is poured into the drum and the drum is rotated

slowly back and forth. Internal partitions divide the samples into equal fractions. The fraction should

be poured again into the drum for further splitting. The process is repeated until a suitable sub-

sample is obtained for counting. The splitter is thoroughly rinsed to recover the organisms, which

should be sticking onto the wall of the drum. The sample is usually splitted into 4 sub-samples. One

of the sub-samples is used for estimation of dry weight, the second for counting the specimens of

common taxa, the third for relative abundance of species and the fourth fraction is kept as reference

collection. Plastic or glass pipettes are also used to take the sub-sample for counting. The Stempel

pipette is used to obtain a certain volume (0.1 to 10 ml). The mesozooplankton sample in a glass

container is diluted to a known volume and is stirred gently. The Stampel pipette is then used to

remove the sub-sample or aliquot for counting” [6].

2.6.1.2 Sample preparation for counting

Several methods can be used to remove organisms from a water sample, such as sedimentation,

centrifugation, and filtration. According to De Bernardi (1984) [18], filtration is the most convenient way

however, Bottrell et al. (1976) [48], suggest that sedimentation is the best method for rotifers, even though

it is practical only at high densities.

✓ One shall note that, with sedimentation it is always laborious and difficult to determine whether all

the organisms have settled. The complete settling of suspended material can take several days. In

addition, removal of the supernatant liquid by siphon requires much care to ensure that animals are

not disturbed [18].



Cautions on filtering (adapted from [18]):

● The filtration must be done at low pressure and as gently as possible to avoid damage to the fragile

organisms or their forced passage through the mesh.

● Rigid filters are preferable because their mesh size is invariable.

✓ Likens & Gilbert (1970) [52] suggest the use of nets with a mesh aperture of 35 µm which,

according to the authors, permits an appropriate retention of even the smaller rotifers.

✓ On the other hand, Schindler (1969) [53] and Ejsmont-Karabin (1978) [54] indicate that

nets with 28 or 10 µm mesh sizes are the most suitable for collecting the smallest forms

especially from nutrient poor mesocosms.

2.6.2 Counting Procedure (Enumeration)

2.6.2.1 Counting chambers

Rotifers and nauplii are counted using either a Sedgewick-Rafter cell or a sedimentation chamber (1-25 ml)

[17]. Sedimentation chambers can be preferred since these allow more flexible volume changes than

Sedgewick-Rafter cells, whereas Makarewicz & Likens (1979) [55] found no difference between counts

obtained using these two chambers.

In addition, rotifers and nauplii can be enumerated in a 1 – 5 ml clear acrylic plastic counting cell fitted with

a glass coverslip [6].

Crustaceans are usually counted using a counting chamber (5 – 10 ml) “with partitions or grooves which

confine the subsample to tracks of a constant width” [49]. The width of the grooves in the tray should be less

than the width of the field at the counting magnification. Petri dish with grid lines on the bottom, a grind

edge and a glass cover to avoid air bubbles, preventing the animals from moving is also usable. Bogorov

design-based counting chambers for mesozooplankton can also be used for counting. An example counting

chamber can be examined in Hydrobios website (REF: https://www.hydrobios.de/produkt/zahlkammer-fur-

mesozooplankton/?lang=de).

A Sedgwick-Rafter cell is not suitable because of size of the crustacean [6].

According to Kaisary and others (2012), an open counting chamber “80 by 50 mm and 2 mm

deep is desirable; however, an open chamber is difficult to move without jarring and disrupting

the count” [6].

✓ As a best practice advice; if the counting chamber do not have partitions or grooves,

mild detergent solution can be placed on the chamber before counting to reduce the

movement of organism

It should be noted that, the chamber for counting the sample may vary with the type of

microscope used.

https://www.hydrobios.de/produkt/zahlkammer-fur-zooplankton/?lang=de

https://www.hydrobios.de/produkt/zahlkammer-fur-zooplankton/?lang=de



2.6.2.2 Counting (enumeration) methodology

✓ “Samples should contain a minimum of 200 animals (for crustaceans: exclusive nauplii) to provide a

good estimate of numbers and species composition. If both crustaceans and rotifers are included in

the analysis, the samples should contain a minimum of 200 animals of the group, which has the

fewest individuals” [1] .

✓ If samples from different dates are pooled, it would be a good idea to use half of the sample for

pooling and keep the rest as a date specific sample.

McCauley explained how to count the rotifer and nauplii as follows: The rotifers and nauplii

“are allowed to settle, approximately 2h per cm height. After settling, the chamber is placed on the

stage of the inverted microscope (i.e. Ütermohl technique) and individuals counted. It is best that

the contents of the whole chamber be enumerated to simplify statistical considerations. This can be

achieved by altering the volume of liquid in the sample and the subsample to bring the number of

organisms in the chamber to the desired level. It is important that all manipulations of sample volume

are recorded and are as accurate as possible so that the original concentration of organisms in the

sample may be calculated” [49].

Counting crustaceans,

“Once the subsample is placed in the chamber, the individuals are counted by moving the chamber

and tallying the individuals encountered in the field of the microscope. Since all of the organisms in

the subsample are usually counted, a homogeneous distribution in the chamber is not needed,

although the volume of the sample should be adjusted so that organisms do not pile up on one

another. Repeated subsamples are then taken until the desired number of individuals have been

counted” [49].

✓ An excel macro including the sheets that can be used for counting individuals of different species

and to make species-area curves (saturation curves) of counted individuals to species is provided

as an attachment to this document (EXCEL 2 – Counting mesozooplankton.xmls). Information on

how to use the excel macro is provided in Appendix II-C.

✓ A high-quality (binocular) dissecting microscope is sufficient for counting Crustacea, although a

light microscope is required for taxonomic identifications [6].

Zooplankton are enumerated in gridded chambers and Bogorov chambers that prevent duplication of

counts. The number of animals counted will vary with the desired degree of sampling precision and the goals

of the study [44].

An example counting (enumeration) procedure was summarized in [31] as follows:

“mesozooplankton is counted using a stereomicroscope until at least 50 individuals of the most

dominant species have been counted [31].



Another counting (enumeration) procedure was summarized in [6] as follows:

“It is recommended that the sub-sample or an aliquot is taken for the common taxa. For enumeration

of mesozooplankton the sub-sample or aliquot of 10 to 25% is usually examined. However, the

percentage of aliquot can be increased or decreased depending on the abundance of

mesozooplankton in the sample” [6].

✓ “For the rare groups, the total counts of the specimens in the samples should be made” [6].

2.6.3 Best Practice Advice on subsampling and counting

✓ Before subsampling, all large organisms such as fish larvae, coelenterates, decapods and other should

be removed and enumerated [6].

✓ It is essential to determine whether subsampling conforms to a random distribution statistically,

since this information is required in determining estimates of counting precision. To confirm that

subsampling is random, replicate counts should be compared with a Poisson distribution using a Chi-

square test ([2] and [49]).

o Lund et al. (1958) [56] recommended that at least 10 sets, each containing > 5 replicate

counts, should be tested for randomness. It is important to test subsampling procedures on

a variety of samples if the density of individual organisms varies considerably among the

samples to be enumerated. If the counts of discrete plankton organisms from a sample are

random and the number of individuals counted is small relative to the total population then,

the counts can be assumed to be distributed according to a Poisson series. If the counts are

not random, then the entire sample should be counted [49]. The entire samples should also

be counted without subsampling in case the mesozooplankton density is low (<200

zooplankter) [6].

✓ The coefficient of variation stabilization technique, that should be utilized to determine population

abundance of a number of taxa simultaneously was summarized in [6] as follows:

“a given sample is split into sub-samples until the 3 most numerous species are present in

numbers of at least 30 animals per sub-sample. All organisms in the sub-sample are then

counted, including all rare species. Alden et al. (1982) supply a table in which the count for

each species in this sub-sample can be looked up, and the sub-sample needed to obtain a

count of at least 30 individuals per species can be read off the chart (i.e., the ½ sub-sample,

the ¼ sub-sample, 1/8, 1/16, 1/32, etc.). Thus, each of the rare species can be counted to the

minimum required precision (i.e. 30 animals per sub-sample) by counting only one more sub-

sample. Counting 30 animals per sub-sample gives a 95% confidence limit of ±30% of the

mean” [6].



● With this technique, the abundance of common species to a predetermined level of

precision can be estimated ([2] and [49]).

● “The coefficient of variation is equal to the inverse of the square root of the number

of individuals counted (CV = 1/√𝑁). For example, if 50 individuals were counted then

the coefficient of variation would be approximately 0.14” [49].

● “This relationship can be used to determine how many individuals should be counted

to obtain a desired level of precision for the estimate of the density of organisms in

a sample” [49].

● Table of calculated values of the coefficient of variation (expressed in fractional

form) of biomass estimates can be used to estimate the number of individuals that

should be counted and to indicate when increases in counting effort are

advantageous. For the table, please visit [49]. For example, moving down a column

at a fixed coefficient of variation indicates the gain in precision with successive

increases of counting effort.

✓ It should be noted that, the calculated values of precision presented in the

mentioned table are based on a number of assumptions and should not be

used without determining whether or not these assumptions are true for

processing a particular sample.

2.6.4 Taxonomy and nomenclature

The list of important studies in identifying mesozooplankton are provided in Appendix II-D.

2.7 Estimating the biomass

✓ larger zooplankters (such as medusae, ctenophores, salps, siphonophores and fish larvae) should be

removed from the mesozooplankton sample and their biomass should be estimated separately. The

total biomass, then, will be the biomass of larger plus the biomass of the rest of the mesozooplankton

[3].

✓ To estimate mesozooplankton biomass, length measurements should be made for minimum 30

rotifers, 30 cladocerans, 30 copepodites and 15 adult copepods per species [31].



2.7.1 Crustacean Zooplankton

2.7.1.1 Measuring the length or dimensions of an organism

Length (or dimensions) of a crustacean mesozooplankton can be measured using an ocular micrometer (a

dissecting microscope equipped with an ocular micrometer) or a computerized measuring system [49].

✓ It should be noted that the use of an ocular micrometer reduces, but does not remove, the possibility

of error in measurements due to variation in the angle and distance between the eye of the observer

and the eyepiece of the microscope [49].

✓ Length of some of the crustacean mesozooplankton is dependent on its developmental stage.

✓ In addition, with the help of digitized plankton images, the analysis can be done using the commonly

used image analysis softwares, such as Zoo/PhytoImage software (licensed under GNU/GPL)

(ZooIMAGE) [57] and ZooScan [58].

For more information on this technique, please refer to [59]–[61].

It should be noted that definition of total length can vary according to the type of the mesozooplankton.

Below are some examples on how to measure the total length:

● The total length of Cladocera can be measured from the top of the head to the point of insertion of

the tail spine. Measurement starting from the eye region to the point of insertion of tail spine is also

used commonly among length measurements of Daphnia [49].

● In case of Copepoda measurements, the length can be measured from “the tip of the cephalothorax

to various points on the urosome, usually excluding the furcal rami” [49].

● McCauley (1984) [49] also noted that, in measuring the length of marine Copepods, maximum width

of the body can be used rather than the length in order to avoid ambiguity in measurements.

✓ For an analysis of the precision of estimates (errors in measurements), please refer to [49].

2.7.1.2 Length-weight relationships (Adopted from Bottrell et al. (1976) [48])

Paterson (2001) claims that the most common method for estimating the dry weight of mesozooplanktonic

Crustacea “relies on converting estimates of length to biomass using length-weight regressions” [2].

✓ As best practice advice, results from measuring systems can be combined with an electronic

database to ease data entry and storage (e.g. Allen et al. (1994) [62]).

An excel sheet, including the length-weight relationships for crustacean mesozooplankton will be provided

as an attachment to this SOP (Excel_1, in preparation). (See Appendix II-E for the list of references that will

be cited).

✓ Length of copepods can be converted into biomass using conversion factors by Paffenhöfer and

Harris (1976) [63], Landry (1983) [64] and Castellani et al. (2005) [65].



2.7.1.3 Estimating the dry weight (Adopted from [49])

1. “To determine the dry weight of crustaceans, individuals are isolated from the sample using a pipette

or a fine dissecting needle (See Furnass & Findley (1975) [66], that presents an apparatus to sort

individuals without picking them up manually, which can be attached to the stage of the dissecting

microscope)

✓ If the weights are for length-weight relationships, then eggs embryos, and gelatinous

sheaths should be removed prior to weighing ([67], [68]).

2. The decision on whether to weigh individuals or groups of animals can be made based on the

expected weight of an individual.

✓ Dumont et al. (1975) [67] suggested that the minimum amount of weighed material

should exceed 5 µg, given that the sensitivity of the most commonly used balances.

3. The animals should be rinsed with distilled water if they are from preserved samples. The animals

are then placed on tared aluminium foil pans or boats ([67], [55]).

✓ The material used as a weighing pan varies among studies. Whatever the material used,

the weight of the pan should be kept to a minimum ([17]).

4. The animals, on their trays, are then placed in an oven and dried for 24-48 h (varied between 2 and

48 h with 24 h being the most frequently used) at 60°C (as in [55], [68]–[72]).

5. After drying, the samples are allowed to cool in a desiccator for 1-2 h and are then transferred to the

balance and weighed using counterbalance techniques to maximize the accuracy of the reading. A

small container of desiccant should be placed in the chamber of the balance to absorb moisture

picked up while transferring the sample.

✓ Placing a container of desiccant inside the housing of the balance and sealing this space

also improves the stability of the reading.

✓ Weighing should be delayed for a short period of time, to allow for stabilization.

To obtain an estimate of precision, replicates of individuals or groups can be weighed. The number

of replicates can then be varied to give a desired level of precision. Please see [73]–[75] for more

details.

2.7.2 Rotifers

2.7.2.1 Predicting dry weight using geometrical formulae (Adopted from [49])

For Rotifers, geometrical formulae are usually combined with appropriate length measures to estimate

biomass.

Estimating the volume of Rotifers, “several different dimensions are measured and then used to calculate a

volume estimate from simple formulae which depend upon the shape of the species (i.e. geometrical

formulae)”.



✓ The geometric formulae provide accurate descriptions of the volume of individuals [49].

✓ A complete description of the technique as well as the list of geometric formulae to calculate the

volumes of over 20 genera can be found in Ruttner-Kolisko (1977) [76] and McCauley (1984) [49] (can

be provided in Excel sheet, Excel_2).

The length of rotifers can be measured using inverted microscope.

✓ The dimensions needed to be used in geometric formulae should be estimated on “slightly

narcotized, living adult specimens, that have not been flattened by a coverslip” [49].

Once the volume has been calculated, it is converted to fresh weight assuming a specific gravity of 1, as the

specific gravity of all species in all habitats equals one [49].

Fresh weights for various Rotifers, which were determined based on the geometric volume calculations, are

presented in Bottrell et al. (1976) [48]. Given fresh weights can be converted to dry weight, as the dry weight

is a constant fraction of fresh weight irrespective of the species being considered, by assuming some constant

value [49].

✓ Constant value was assumed as 0.05 (dry:wet) in [77] and 0.1 (dry:wet) in [70] and [78].

Measuring the dry weight by microbalance or weighing is impractical and inefficient for rotifers. As the small

size of rotifers makes it impossible to weigh individual animals of most species [49]. They should be weighed

as a group and a mean weight should be determined by accounting for the number of individuals used [49].

✓ Ruttner – Kolisko (1977) [76] reminded that this technique requires more time and less efficient in

determining weight than volume and weight determination with geometric formulae.

If considered, [55] and [67] described this procedure clearly. It is also summarized by [49].

2.7.2.2 Estimating Mass as Carbon Content (Adopted from [49])

The technique proposed by Latja & Salonen (1978) [79], to determine the carbon content of individuals

“requires high temperature combustion of the sample and measurement of the carbon dioxide evolved using

an infra-red gas analyzer. It requires 10000 times less material and processes samples 20 times faster than

current methods of calorimetry” [49].

“Using this technique, sub-microgram quantities of carbon can be measured. This sensitivity is considerably

better than the best microbalance, and the carbon content of an individual animal as small as a rotifer can

be determined in < 1 minute” (as in [79] and [49]).

The technique was summarized by [49] as follows:

1. “An individual is transferred to the combustion tube using forceps, and then combusted at 950°C.

✓ If the animal is transferred with a pipette, then the carbon content of the volume of water

used must be determined and subtracted from the weight of the animal.



2. The carbon dioxide produced is measured using an infra-red gas analyzer. A complete description of

the apparatus, its calibration, recovery statistics, and design for constructing the quartz combustion

tube can be found in Salonen (1979) [80]” [49].

3. The probable conversion factor (µg C per µg dry weight) for different taxa can also be used and can

be reviewed from [81] and [82].

2.7.3 Estimating biomass: other methods in literature

The biomass can also be estimated by the following methods (as provided in [6]):

● Volumetric (displacement volume and settling volume) method (in the field or laboratory): The

total mesozooplankton volume is determined by the displacement volume method. In this

method;

a) “the mesozooplankton sample is filtered through a piece of clean, dried netting material.

b) the mesh size of netting material should be the same or smaller than the mesh size of the net

used for collecting the samples.

c) The interstitial water between the organisms is removed with the blotting paper.

✓ While blotting, due care should be taken not to exert too much pressure as

to damage the delicate organisms or specimens.

d) The filtered mesozooplankton is then transferred with a spatula to a measuring cylinder with

a known volume of 4 % buffered formalin.

e) The displacement volume is obtained by recording the volume of fixative in the measuring jar

displaced by the mesozooplankton.

f) The settled volume is obtained by making the sample to a known volume in the measuring jar”

[6].

✓ Using this technique, the plankton is allowed to settle for at least 24 hours

before recording the settled volume [6].

● Gravimetric (wet weight, dry weight and ash free dry weight) method (preferably in laboratory):

In this method:

a) “Filter the mesozooplankton and remove the interstitial water by a blotting paper.

✓ While blotting, due care should be taken not to exert too much pressure as

to damage the delicate organisms or specimens.

b) The mesozooplankton weight is taken on predetermined or weighed filter paper or aluminium

foil.

Express the net weight in grams”.

c) The dry weight method is dependable as the values indicate the organic content of the

plankton. Analysis such as the dry weight is determined by drying an aliquot of the

mesozooplankton sample in an electric oven at a constant temperature of 60ºC.

✓ The whole or total sample should not be dried because subsequent analyses

such as enumeration of common taxa and identification of the species wouldn’t

be possible after drying the sample.



d) The dried aliquot is kept in a desiccator until weighing. Express the values in milligram.

e) Ash free dry weight method is also occasionally used for biomass estimation” [6].

✓ This method can be used for estimating the biomass of gelatinous

mesozooplankton.

2.7.4 Size Distribution

Differences in mesozooplankton size distribution can be assessed by calculating size diversity and normalised

biomass-size spectrum (NSS).

2.7.5 Size Diversity

Size diversity can be calculated by using “function of probability density of individuals with respect to size”

[83] as described in [84], [85]. As it is a continuous function, this approach provides the advantage of avoiding

“the arbitrariness introduced when using size classes” [83].

Moreover, Quintana and colleagues (2008) provided the methodology, which is based on the Shannon-

Wiener diversity expression [86] and adapted to a continuous variable, such as the body size; to estimate the

size diversity by using the individual size measurements [87].

● One should note that, if size diversity is high, whether the size range (i.e. the range representing the

minimum and maximum length of species in µm) is wider and/or there are similar proportions of the

different sizes [83], [87]–[89].

2.7.6 Normalised biomass-size spectrum (NSS)

As stated by Brucet and colleagues (2010), “biomass-size spectrum describes how biomass of organisms is

distributed along size classes” [90]. One should see the book by Kerr and Dickie, published in 2001, ‘The

Biomass Spectrum: A Predator-Prey Theory of Aquatic Production’ [91] for more details on how to assess the

size distribution with NSS approach.

As a best practice advise, using size diversity approach is advantageous over NSS as the former one does not

require statistical analysis (in contrast to NSS) and “is represented by a single value” [92].

2.8 Quality assurance and quality control

According to [93], systematic or random errors can occur during the sampling operations. Systematic errors

are generally due to the “poor sampling practices or equipment design failures” which are usually constant

[93]. On the other hand, random errors are generally unavoidable or unpredictable. One shall follow an

effective Quality Assurance/Quality Control (QA/QC) strategy during the experiment to identify, quantify and

control the errors. Standardization of sampling and analysis methods, taking replicate samples and analyses,



or following a laboratory accreditation scheme are some examples for QA practices that can be followed

during the experiment. Quality Assurance (QA) strategies are defined by the scientist-in-charge to ensure

that the sample data meets Data Quality Objectives (DQO), which shall be defined prior to sampling. On the

other hand, Quality Control (QC) is “the system of guidelines, procedures and practices designed to regulate

and control the quality of products and services, ensuring they meet pre-established performance criteria

and standards”. The QC practices that can be followed are as follows: taking “sample blanks, replicates,

splits”, having and following “equipment calibration standards”, determination of “sample container size,

quality, use and preservative amount” prior to sampling.

The WISER project results highlighted the information below for QA of sampling and counting of plankton:

“1) Details of microscopes, chambers (individually identified and calibrated) and calibration of all

ocular/objective combinations should be recorded in a note book and kept for reference. If fixed

volume pipettes are used, these should be calibrated annually.

2) Checks for random distribution of sample should be done visually at low magnification for each

sample. Some simple checks include:

Comparing the number of observations in:

(a) half a chamber with the other half

(b) comparing counts in the 1st transect with the 2nd transect

(c) comparing counts in the first 20 field of view with the next 20 fields.

A more detailed check using simple Chi squared test should be done if a sample does not appear to

be randomly sedimented or 1 sample every 3 months or so” [94].

According to the ‘Common Implementation Strategy for the Water Framework Directive (2000/60/EC)’, the

intercalibration exercises in between laboratories will provide a “continuous quality assurance system”, by

ensuring the results meet targeted levels [95]. In order to perform intercalibration in the AQUACOSM

community, there shall be QA measures in each of the mesocosm facility. The common QA measures, that

are determined based on this SOP and the valid sampling and analysis methods used during the experiments,

that shall be taken by each mesocosm facility are listed as follows:

● “Field sampling and sample label

● Sample storage and preservation;

● Laboratory analysis” [95].

Further suggestions to assure sampling quality (not specific to mesozooplankton analysis) are as follows:

● One shall establish of a “routine internal quality control” and participate in “external quality

assurance (QA) schemes” [95].

● In addition, one shall consider the resolution/degree of identification in counting and take

pictures of unidentified species or species which have unsure identification.

For detailed QA methodology provided for sampling, analysis and counting of mesozooplankton, please visit

EN 14996:2006 [96] and EN 15110:2006 [1] .



2.9 References 2 - Zooplankton

[1] CEN, “EN 15110: 2006: Water Quality - Guidance

standard for the sampling of zooplankton from

standing waters,” 2006.

[2] M. J. Paterson, “Ecological Monitoring and

Assessment Network (EMAN) Protocols for

measuring biodiversity: Zooplankton in fres waters,”

2001.

[3] S. Goswami, “Zooplankton Methodology, Collection

& identyification - A field manual,” Natl. Inst.

Oceanogr., p. 16, 2004.

[4] The Editors of Encyclopaedia Britannica, “Halocline |

Oceanography | Britannica.com,” Encyclopædia

Britannica, inc., 1998. [Online]. Available:

https://www.britannica.com/science/halocline.

[Accessed: 05-May-2019].

[5] US EPA, “Standard Operating Procedure for

Zooplankton Sample Collection and Preservation

and Secchi Depth Measurement Field Procedures -

LG402,” 2013.

[6] S. Kaisary et al., “Coastal Water Quality

Measurements Protocol for COMAPS Programme,”

pp. 1–110, 2012.

[7] WHO, Handbook: Good Laboratory Practice (GLP):

Quality practices for regulated non-clinical research

and development, 2nd ed. Switzerland: TDR/WHO,

2009.



[9] Council Directive, “89/391/EEC of 12 June 1989 on

the introduction of measures to encourage

improvements in the safety and health of workers at

work,” Off. J. Eur. Communities , vol. 183, no. 29, p.

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[10] OSHA, “European directives on safety and health at

work - Safety and health at work,” 2017. [Online].

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16-Jun-2017].

[11] Division on Earth and Life Studies Committee on

Prudent Practices in the Laboratory Board on

Chemical Sciences and Technology, Prudent

practices in the laboratory : handling and

management of chemical hazards, Updated Ve.

Washington, D.C.: National Academy of Sciences,

2011.

[12] ECHA, “Formaldehyde - Substance Information.”


https://echa.europa.eu/substance-information/-

/substanceinfo/100.000.002. [Accessed: 16-Jun-

2017].

[13] Ministerium Für Ein Lebenswertes Österreich,

“LEITFADEN ZUR ERHEBUNG DER BIOLOGISCHEN

QUALITÄTSELEMENTE TEIL C - ARBEITSSICHERHEIT,”

Wien, 2016.










pursuant to Article 1(4) of ,” Off. J. Eur. Communities,

vol. 69, 2000.

[16] P. S. Welch, “Limnological methods,” 1948.

[17] W. T. Edmondson and G. G. Vinberg, “Manual on

methods for the assessment of secondary

productivity in fresh waters,” 1971.

[18] R. De Bernardi, “Methods for the estimation of

zooplankton abundance,” in A Manual on Methods

for the Assessment of Secondary Productivity in Fresh

Waters, 1984, pp. 59–86.

[19] D. R. DeVries and R. A. Stein, “Comparison of three

zooplankton samplers: a taxon-specific assessment,”

J. Plankton Res., vol. 13, no. 1, pp. 53–59, 1991.

[20] R. Harris, P. Wiebe, J. Lenz, H.-R. Skjoldal, and M.

Huntley, ICES zooplankton methodology manual.

Academic Press, 2000.

[21] J. Schwoerbel, Methods of hydrobiology:(freshwater

biology). Elsevier, 2016.



[22] N. Manickam, P. Santhanam, and P. Saravana

Bhavan, “Techniques in the Collection, Preservation

and Morphological Identification of Freshwater

Zooplankton,” in Basic and Applied Zooplankton

Biology, Singapore: Springer Singapore, 2019, pp.

139–195.

[23] P. Santhanam, P. Pachiappan, and A. Begum, “A

Method of Collection, Preservation and

Identification of Marine Zooplankton,” in Basic and

Applied Zooplankton Biology, Singapore: Springer

Singapore, 2019, pp. 1–44.

[24] HYDRO-BIOS, “HYDRO-BIOS,” 2017. [Online].

Available: https://www.hydrobios.de/.

[25] G. L. Clarke and D. F. Bumpus, “The Plankton Sampler

- an instrument for quantitative plankton

investigations,” Linnological Soc. Am., vol. Special

Pu, no. 5, pp. 1–8, 1940.

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Co-funded by the European Union D4.1 Standard Operating Procedures| 3 Microbial Plankton


3. Microbial Plankton


Table 3-1: Main terms and abbreviations that will be used throughout the text

Plankton Organisms drifting or suspended in water, consisting chiefly of minute plants

or animals, but including larger forms having only weak powers of locomotion.

Biomass The amount of living matter present in the plankton sample.

BCG Bacterioplankton community growth. Measured by uptake of radioactively

labelled substrates like the DNA base thymidine or the amino acid leucine.

TCF Thymidine conversion factor from mol thymidine assimilated to cells produced.

Determined by simultaneous measurement of thymidine uptake and cell

production by microscopy in seawater cultures. Alternatively, calculated from

reported thymidine content in the average bacterial cell.

LCF Leucine conversion factor. Determined as for TCF but using the amino acid

leucine.

Flow cytometry Flow cytometry is a technology used to analyse the physical and chemical

characteristics of particles in a fluid as it passes through at least one laser. Cells

have natural fluorescence or are often labelled with fluorescent markers so

that light is first absorbed and then emitted in a band of wavelengths. The

fluorescence as well as other characteristics of single particles such as their

relative granularity, and its internal complexity can be quickly examined (up to

thousands of cells per second) and the data gathered are processed by a

computer.

Viruses A small infective agent that typically consists of a nucleic acid molecule in a

protein coat is too small to be seen by light microscopy that is able to multiply

only within the living cells of a host organism. Viruses can infect all types of life

forms, from animals and plants to microorganisms, including bacteria and

archaea.

Prokaryotes Usually unicellular organisms, sometimes multi cellular organisms, that lack a

distinct nucleus, mitochondria, or any other membrane-bound organelle due

to the absence of internal membranes.

https://en.wikipedia.org/wiki/Prokaryote - cite_note-NCSU-1 The word

prokaryote comes from the Greek πρό (pro) "before" and κάρυον (karyon) "nut

or kernel". Bacteria are among the best-known prokaryotic organisms.

https://en.wikipedia.org/wiki/Life_forms

https://en.wikipedia.org/wiki/Life_forms

https://en.wikipedia.org/wiki/Microorganism

https://en.wikipedia.org/wiki/Bacteria

https://en.wikipedia.org/wiki/Archaea

https://en.wikipedia.org/wiki/Unicellular_organism

https://en.wikipedia.org/wiki/Cell_nucleus

https://en.wikipedia.org/wiki/Mitochondrion

https://en.wikipedia.org/wiki/Organelle

https://en.wikipedia.org/wiki/Prokaryote%20-%20cite_note-NCSU-1

https://en.wikipedia.org/wiki/Greek_language

https://en.wikipedia.org/wiki/Nut_(fruit)

https://en.wikipedia.org/wiki/Seed



Eukaryotes Any cell or organism that possesses a clearly defined nucleus. The eukaryotic

cell has a nuclear membrane that surrounds the nucleus, in which the well-

defined chromosomes (bodies containing the hereditary material) are located.

Eukaryotic cells also contain organelles, including mitochondria, a Golgi

apparatus, an endoplasmic reticulum, and lysosomes. Their name comes from

the Greek εὖ (eu, "well" or "true") and κάρυον (karyon, "nut" or "kernel")

eukaryotes may also be multi-cellular and include organisms consisting of many

cell types forming different kinds of tissue. Animals, plants and fungi are the

most familiar eukaryotes.

Protists Any eukaryotic organism that is not an animal, plant or fungus. While

exceptions exist, they are primarily microscopic and unicellular, or made up of

a single cell. The cells of protists are highly organized with a nucleus and

specialized cellular machinery called organelles.

A general description for water sampling will be covered under the Water Chemistry SOP (Version 2.0).


Table 3-2: Reagents and recipes used in different methods


Acid Lugol’s Iodine KI, I2, glacial acetic acid, distilled water

Put 100 gr KI in 1L distilled water, add 50 gr iodine and

100 mL glacial acetic acid.

Fume

Cabinet/Hood

Formaldehyde (37% by volume) Fume

Cabinet/Hood

Ethanol (96% or 99%) Ethanol, water

Instead of pure ethanol, cheaper methylated spirit can

be used. If zooplankton is stored for molecular analysis,

a solution from pure ethanol should be used (70%).

Solvent

cabinet

Buffered sucrose formation (Buffered

Formalin)

Glutaraldehyde (25%, for electron

microscopy quality)

Fridge (4°C)

https://www.britannica.com/science/cell-biology

https://www.britannica.com/science/nucleus-biology

https://www.britannica.com/science/nuclear-membrane

https://www.britannica.com/science/chromosome

https://www.britannica.com/science/organelle

https://www.britannica.com/science/mitochondrion

https://www.britannica.com/science/Golgi-apparatus

https://www.britannica.com/science/Golgi-apparatus

https://www.britannica.com/science/endoplasmic-reticulum

https://www.britannica.com/science/lysosome

https://en.wikipedia.org/wiki/Greek_language

https://en.wiktionary.org/wiki/%CE%B5%E1%BD%96

https://en.wiktionary.org/wiki/%CE%BA%CE%AC%CF%81%CF%85%CE%BF%CE%BD

https://en.wikipedia.org/wiki/Multicellular_organism

https://en.wikipedia.org/wiki/Cell_type

https://en.wikipedia.org/wiki/Tissue_(biology)

https://en.wikipedia.org/wiki/Eukaryotic

https://en.wikipedia.org/wiki/Organism

https://en.wikipedia.org/wiki/Animal

https://en.wikipedia.org/wiki/Plant

https://en.wikipedia.org/wiki/Fungus



SYBR Green I (other stains) SYBRGreen I (ThermoFisher , S7563 / S7567 / S7585) is

supplied as a 10,000X concentrate in DMSO. Store at <-

20°C. Avoid direct light and several freeze/thaw cycles.

For FCM, Prepare working solutions, of 200x for bacteria

and 50x concentration for viruses by diluting in DMSO or

Tris-EDTA. The working solutions should be stored in

dark at 4°C for short periods (ca. a week) or at -20°C for

long periods.

For viral analysis on microscope, prepare a 25x working

solution.

Freezer (-

20°C)

Paraformaldehyde (PFA) solution (2%

final concentration in the sample)

See appendix for detailed information about preparing

PFA solution.

Freezer (-

20°C)

p-phenylenediamine ant-fade

mounting solution

Make a 10% (wt/vol) solution by weight with 0.02 mm

filter- autoclaved dH2O.

Freezer (-

20°C)

Acridine orange Acridine orange (Sigma artikelnr. A6014-10G, clear red

quality) is mixed with MilliQ-water to 3 mg mL-1. The

solution is filtered through a 0.2 µm sterile filter (e.g.

Acrodisc) into the filtration funnel.

Fridge (4°C)

maximum 8

weeks

Immersion oil, low fluorescent For example, Type A, formula code 1248. Cat. No. 16482

(R.P. Cargille laboratories, Inc. Cedar Grove, N.J. 007009,

USA).

Room

temperature

(4-40°C)

Fluorescent beads, standard Duke Scientific, Polymere Microspheres™ Green

Fluorescing, 1% solids, 1,4*1011 kulor mL-1, diameter

0.519 µm, CV<5%, Cat. No. G500) används som standard

materiel. Volume is 0.073 µm3 (0.063-0.085 µm3). Make

a working solution of 1x106 beads mL-1

Fridge (4°C)

TCA, Trichloracetic acid

3.3 Health and Safety – Safe Disposal

Please see the SOPs: Water Chemistry (section 5Water Chemistry) and Phytoplankton (section 1) for general

health and safety instructions regarding sampling and general lab procedures and best practices.

For specific hazardous materials and reagents mentioned in this document, such as SYBRGreen I,

Glutaraldehyde, PFA, etc. please, refer to the appropriate COSHH forms by the manufacturer regarding

handling and safe disposal.

Always follow the safety instructions and risk assessment protocols of the lab.



3.4 Sample Integrity – Sampling – Best Practices

3.4.1 Methods

3.4.1.1 Viruses

It is recommended to use Flow Cytometry (FCM) for the counting of viruses due to the small size of viral

like particles and the photobleaching of the DNA stains that are used. However, an Epifluorescence

microscope can be used if a flow cytometer is not available.

i) Microscopy

Fix samples with glutaraldehyde (0.5% final concentration) for approximately 30 min at 4°C. Prepare the

microscope slides within 4 h after fixation.

If it is not possible to prepare the slides within 4 h after fixation, flash freeze the samples in liquid nitrogen

and store at -80°C until slide preparation.

Prepare the appropriate working solutions of SYBRGreen I and p-phenylenediamine anti-fade mounting

medium.

Gently thaw the samples (if have been frozen).

Use 0.02 μm Anodisc or polycarbonate filters to filter the appropriate volume of the sample.

After filtration, dry the filters completely either by gently rubbing against a Kimi wipe or using a heating

block.

Stain the filters with SYBRGreen I, dry them again and then place them on a microscopy slide. Place the

appropriate amount of the p-phenylenediamine anti-fading mounting medium and put the slide cover.

To view the slide and count the viruses, use a 100x fluorescence oil-immersion objective with immersion

oil. SYBR Green I binds to dsDNA and is excited with a maximum at 488 nm. A wide BP blue-excitation

filter and long-pass (LP) green-emission filter will optimize observation of the cells and viruses.

A detailed protocol on viral and prokaryotic cells analysis with epifluorescence microscopy has been

published by Patel et al. (2007).

ii) FCM

Fix samples with glutaraldehyde (0.5% final concentration) for approximately 30 min at 4°C. Flash freeze

the samples in liquid nitrogen and store at -80°C until analysis.

Before analysis, thaw samples gently and dilute to appropriate concentration for the instrument (for

example, less than 700 events/sec for BD FACSCalibur) using Tris-EDTA buffer at pH 8.0.

Stain with SYBRGreen I (e.g. SYBRGreen I, ThermoFisher, S7563) to a final dilution of 5 x 10-5 of the

commercial solution (e.g. use 5 μL of the 50x working dilution – mentioned at Table 3-2– at 495 μL of



diluted sample). Incubate at 80°C for 10 min in the dark. Allow sample to cool down for 5 min prior to

analysis.

Use a laser providing blue light (488 nm) and set trigger on green fluorescence. Group, detect and

enumerate various groups of viruses based on difference in green fluorescence. Most of the times, viruses

are separated in three groups based on their fluorescence.

For more information regarding the viral analysis on FCM refer to Brussaard et al. (2004) and Mojika et al.

(2014).

3.4.1.2 Heterotrophic Bacteria

iii) Microscopy

Fix samples with 37% formaldehyde. A microscopic slide should be prepared within 7 days. Microscopic

slides can be stored at -20°C for 70 days before analysis. Apply a GF/C filter on a filter support of a

multifilter unit wetted with MilliQ water. Place a black 0.2 µm polycarbonate filter with shiny side up on

top of the GF/C filter. Additional filter pore sizes may also be appropriate, i.e. 3.0 µm for discriminating

amongst particle-associated and free-living bacteria (Hobbie et al., Azam et al), particularly when algal or

zooplankton densities are high. Apply vacuum to make the filters stick to the support. Shake the sample

for 10 s and take out the appropriate volume of sample for a proper density of cells (the volume depends

on the trophic status of the environment). Wash the tip in MilliQ water between transfers. Filter the

sample dry with -13 kPa vacuum. Remove vacuum. Add 15 drops of Acridine orange (AO) stain (3 mg mL-

1) or l−1 DAPI (4,6-diamidino-2-phenylindole) through a 0.2 µm filter (e.g. Acrodisc). Incubate for 10 min,

label the glass slide in the meantime. Filter until surface is dry. Remove vacuum. Wash with 1 mL MilliQ

water, filter dry. Dry filter in the air holding it with forceps for 45 s. Mount on glass slide with immersion

oil spread on a spot. Add a drop of immersion oil on top of the filter and apply a cover slip. Analyse the

sample in an epifluorescence microscope with filter set (FS 09, 450-490, FT 510, LP 520) and 63x Plan-

Apochromat objective (Hobbie et al. 1977). Preferably take 5 images per sample with a high resolution

black and white camera. Count cell numbers, morphology and size by a neural network analysis software

like LabMicrobe (BioRas, Denmark, Blackburn et al. 1998). Manual counting using ocular square grids like

the Miller Square grid can also be used. The magnification factor between counted area and total filtered

areas needs to be determined for each method. Fluorescent beads are used to calibrate the system

regarding particles counted and their size. The cell morphologies of cocci, rods and vibroids in the size

range 0.2-2 µm are counted as bacteria. Sample images of these are used to train the neural network. Cell

volumes can be used to calculate bacterial carbon density and biomass (Norland 1993).

iv) FCM

Fix samples with glutaraldehyde (0.5% final concentration) for ca 30 min at 4°C. Flash freeze the samples

in liquid nitrogen and store at -80°C until analysis. Before analysis, thaw samples gently, dilute to

appropriate concentration and stain with SYBRGreen I (e.g. MolecularProbes, Eugene, Oregon, USA) for

10 min in room temperature (Marie et al. 1999). Use a laser providing blue light (488 nm) and set trigger



on green fluorescence. Group, detect and enumerate various groups of bacteria based on difference in

green fluorescence and scatter properties.

3.4.1.3 Autotrophic bacteria and nanophytoplankton:

v) Microscopy

For the enumeration of autotrophic bacteria (Synechococcus spp.) and nanoflagellates, samples are fixed

with 1.8% formaldehyde buffered with sodium tetraborate decahydrate and filtered for particle removal

through 0.45 μm filters. Fixation is allowed for 1 h in the dark and at 4°C. Subsequently, samples are stained

with 0.2 mg l−1 DAPI (4,6-diamidino-2-phenylindole) for 10 min and filtered through 0.6 μm black

polycarbonate membranes (Porter and Feig 1980). Filters are mounted on glass slides and stored at -20°C

until analysis. Enumeration is performed using epifluorescence microscopy under UV-excitation. Autotrophic

bacteria and nanoflagellates are distinguished by their orange and red fluorescence, respectively, observed

under blue light excitation; at least 50 fields are counted for each sample. Nanoflagellates are separated into

four size classes (< 2 μm, 2-5 μm, 5-10 μm and >10 μm) using an ocular micrometer. An ellipsoid shape is

assumed for nanoflagellates and the biovolume is calculated separately for each size class using the

measured dimensions. Nanoflagellate biomass is calculated using the conversion factor 183 fg C μm-3

described by Caron et al. (1995). Synechococcus abundance data are converted into C biomass using 250 fg

C cell-1 (Kana and Glibert, 1987).

vi) FCM

Phytoplankton counts are best obtained from fresh, but can also be obtained from fixed (glutaraldehyde)

samples.

Use a laser providing blue light (488 nm), set trigger on red fluorescence and group, detect and enumerate

various groups of phytoplankton (typically Synechococcus, Prochlorococcus, picoeukaryotes,

nanoeukaryotes, cryptophytes) based on chlorophyll (red fluorescence) and phycoerythrine

autofluorescence and side-scatter signal (SSC) signals (e.g. Larsen et al. 2001).

3.4.1.4 Heterotrophic Flagellates

vii) Microscopy

For the enumeration of heterotrophic nanoflagellates, samples are fixed with 1.8% formaldehyde

buffered with sodium tetraborate decahydrate and filtered for particle removal through 0.45 μm filters.

Fixation is allowed for 1 h in the dark and at 4°C. Subsequently, samples are stained with 0.2 mg l−1 DAPI

(4,6-diamidino-2-phenylindole) for 10 min and filtered through 0.6 μm black polycarbonate membranes

(Porter and Feig 1980). Filters are mounted on glass slides and stored at -20°C until analysis. Enumeration

is performed using epifluorescence microscopy under UV-excitation. Autotrophic nanoflagellates are

distinguished from heterotrophic ones by the red fluorescence of their chloroplasts, observed under blue

light excitation; at least 50 fields are counted for each sample. Nanoflagellates are separated into four

size classes (<2 μm, 2-5 μm, 5-10 μm and >10 μm) using an ocular micrometer. An ellipsoid shape is



assumed for nanoflagellates and the biovolume is calculated separately for each size class using the

measured dimensions. Biomass is calculated using the conversion factor 183 fg C μm-3 described by Caron

et al. (1995).

viii) FCM

Fix samples with glutaraldehyde (0.5% final concentration) or paraformaldehyde (1% final concentration)

for at least 2 h at 4°C. Flash freeze the samples in liquid nitrogen and store at -80°C until analysis. Thaw

samples gently before analysis and stain with SYBRGreen I (e.g. MolecularProbes, Eugene, Oregon, USA)

for at least 10 min in room temperature (Zubkov et al. 2007). Using a laser providing blue light (488 nm),

set trigger on green fluorescence and discriminate HNF population(s) from nano-sized phytoplankton

based on green vs. red fluorescence and from large bacteria on plots of side scatter vs. green fluorescence

following the recommendations of Christaki et al. (2011).

3.4.1.5 Micro-Plankton (Dinoflagellates, Ciliates)

For ciliate and dinoflagellate enumeration, samples (the sample volume depends on the trophic status of

the environment) are preserved with borax-buffered formaldehyde (final concentration 2%) or with acid

Lugol’s solution (final concentration 2%). The samples are stored at 4°C in the dark and examined within

3 mo of collection. Before examination, samples are left to settle for about 24 h in Utermöhl chambers

(Utermöhl 1931) and are finally examined with an inverted microscope at 200×. The microscope may be

equipped for transmitted light, phase-contrast and epifluorescence.



Figure 3-1: (A, B) Biparametric flow cytometry plots with the applied grouping of the different phytoplankton groups indicated by the solid line. Groups are coloured according to their grouping in order to appear on both (A) BL2 (orange fluorescence) vs. SSC (side scatter) and (B) BL3 (red fluorescence) vs. BL2. (C) Microscope image showing the difference in the autofluorescence spectrum of Synechococcus (appear orange) and pico-eukaryotes (appear green). The long orange cell is a diatom.

Blue light excitation (DM 500 nm dichroic mirror, BP 420 to 480 nm exciter filter, BA 515 nm barrier filter

and a 100 W mercury burner) is used to detect chlorophyll autofluorescence and to distinguish plastidic

from non-plastidic ciliates. A problem associated with the preservative choice is the possibility of affecting

the apparent importance of loricate and aloricate ciliates since tintinnids are expected to be more robust

to preservation. Buffered formaldehyde (final concentration 2%) is often used especially when we need

to know about the trophic status of ciliates. Stoecker et al. (1989) established that samples preserved in

buffered formaldehyde lost 10 to 20% of aloricate ciliates compared to samples preserved in acid Lugol’s

iodine solution, whereas Revelante & Gilmartin (1983) estimated this loss to be 30 to 70%.

3.4.2 Processes

1) Bacterioplankton community growth (Smith and Azam 1992)

Wash a 50 mL polypropylene tube (e.g. Falcon™) with sample water and then collect the sample. Fill 2-4

micro vials (polypropylene, e.g. Eppendorf) with 1 ml sample water. Where many true replicates occur,

1 sample and one or fewer controls can be used. Otherwise, 3 replicate samples and one control are

recommended. Wash the tip with MilliQ water and sample water between each sample type. Add 100

µL ice cold 50% trichloroacetic acid (TCA) to the control samples. Vortex 3 s and invert the tube to mix.



Incubate 5 min at 2°C. Take out the total required amount of 3H-Thymidine (80 000 Ci mol-1,

concentration of 12.5 µM, 1mCi mL-1) to a micro vial. Transfer 2 µL of the isotope (20 nmol dm-3 final

conc.) to the wall above the liquid to each vial starting with the samples and finish with the controls. Mix

all vials 3 s and note the time for incubation start. Incubate in a thermos with the in-situ temperature for

1 hour. Stop the incubation by putting the vials into a micro vial cooling-block (2°C, e.g. Eppendorf

Thermostat C for 5 min.). Note the stopping time. Add 100 µL of ice cold 50% TCA to the sample tubes

only. Incubate for 5 min. Samples can be stored at this temperature up to 7 days. Centrifuge the samples

with necks facing outwards at 16000 g (13000rpm) for 10 min. Remove the supernatant and any

moisture in the lid or walls by a tap water vacuum evaporator with a drawn Pasteur glass pipette. Add 1

ml of 5% TCA, turn the vial upside down removing any air bubble and then vortex 5 s. Centrifuge again

as above. Remove the supernatant as above. Finally add 1 mL Scintillation liquid (e.g. Optiphase HiSafe)

to each vial. Place in scintillation vials (5 mL size). Samples can be stored at room temperature in the dark

for at least 1 month. Count samples in the tritium channel in a calibrated scintillation counter with applied

quench correction. Calculate the uptake of thymidine according to the reference above. Convert to cell

production by using empirical or theoretical thymidine conversion factor (Wikner and Hagström 1999).

Calculation to biomass production can be done by multiplying with the per cell carbon content from

bacterial biomass estimates. When bacterial biomass production is wanted 3H-leucine (Perkin-Elmer Life

Science Products; 170 mCi mmol-1) may be used instead of thymidine, applying the theoretical conversion

factor for leucine (del Giorgio et al. 2011).



3.5 References 3 – Microbial Plankton

Azam, F., Holm-Hansen, O. Use of tritiated substrates in the

study of heterotrophy in seawater. Mar. Biol. 23, 191–

196 (1973). https://doi.org/10.1007/BF00389484

Bell R.T. (1993) Estimating production of heterotrophic

bacterioplankton via incorporation of tritiated

thymidine. In Kemp P.F., B.F. Sherr, E.B. Sherr and J.J.

Cole [eds.], Handbook of methods in aquatic

microbial ecology. Lewis Publishers: p. 495-503.

Blackburn N. et al. (1998) Rapid Determination of Bacterial

Abundance, Biovolume, Morphology, and Growth by

neural Network-Based Image Analysis. Appl. Environ.

Microbiol. 64(9): 3246-3255.

Brussaard C.P.D. (2004) Optimization of Procedures for

Counting Viruses by Flow Cytometry. Appl. Env.

Microbl. 70: 1506-1513.

Caron D.A., Dam H.G., Kremer P., Lessard E.J., Madin L.P.,

Malone T.C., Napp J.M., Peele E.R., Roman M.R.,

Youngbluth M.J. (1995) The contribution of

microorganisms to particulate carbon and nitrogen in

surface waters of the Sargasso Sea near Bermuda.

Deep-Sea Res. Part I 42: 943-972.

https://doi.org/10.1016/0967-0637(95)00027-4.

Christaki U., Courties C., Massana R., Catala P., Lebaron P.,

Gasol J.M. et al. (2011).Optimized routine flow

cytometric enumeration of heterotrophic flagellates

using SYBR Green I. Limnol. Oceanogr. Methods 9:

329-339. doi: 10.4319/lom.2011.9.329.

Fuhrman J.A. and Azam F. (1982) Thymidine incorporation as

a measure of heterotrophic bacterioplankton

production in marine surface waters: Evaluation and

field results. Mar. Biol. 66: 109-120.

Hobbie J.E., Daley R.J. and S. Jasper (1977) Use of

nucleporefilters for counting bacteria by fluorescence

microscopy. Appl. Environ. Microbiol. 33: 1225-1228.

Kana T., Glibert P.M. (1987) Effect of irradiances up to 2000

mE m-2 s-1 on marine Synechococcus WH 7803-I.

Growth, pigmentation and cell composition. Deep Sea

Research 34: 479-516.

Larsen A., Castberg T., Sandaa R.A., Brussaard C.P.D. Egge J.,

Heldal M. et al. (2001). Population dynamics and

diversity of phytoplankton, bacteria and viruses in a

seawater enclosure. Mar. Ecol. Prog. Ser. 221: 47-57.

doi:10.3354/meps221047.

Marie D., Partensky F., Jacquet S. and Vaulot D. (1997).

Enumeration and cell cycle analysis of natural

populations of marine picoplankton by flow

cytometry using the nucleic acid stain SYBR Green I.

Appl. Environ. Microbiol. 63: 186-93.

Mojika K.D.A., Evans C., Brussaard C.P.D. (2014) Flow

cytometric enumeration of marine viral populations

at low abundances. Aquat. Microb. Ecol. 71: 203-209.

Norland S. (1993) The relationship between biomass and

volume of bacteria. In Kemp P.F., B.F. Sherr, E.B. Sherr

and J.J. Cole [eds.], Handbook of methods in aquatic

microbial ecology. Lewis Publishers: p. 303-307.

Patel A., Noble R.T., Steele J.A., Schwalbach M.S., Hewson I.,

Fuhrman J.A. (2007) Virus and prokaryote

enumeration from planktonic aquatic environments

by epifluorescence microscopy with SYBR Green I.

Nat. Protoc. 2: c269-276.

Porter K.G., Feig Y.S. (1980) The use of DAPI for identifying

and counting aquatic microflora. Limnol. Oceanogr.

25: 943-948.

https://doi.org/10.4319/lo.1980.25.5.0943.

Revelante N. and Gilmartin M. (1983) Microzooplankton

distribution in the Northern Adriatic Sea with

emphasis on the relative abundance of ciliated

protozoans. Oceanol Acta 6: 407-415.

Simon M. and Azam F. (1989) Protein content and protein

synthesis rates of planktonic marine bacteria. Mar.

Ecol. Prog. Ser. 51: 201-213.

Smith D.C. and Azam F. (1992) A simple, economical method

for measuring bacterial protein synthesis rates in

seawater using 3H-leucine. Mar. Microb. Foodwebs 6:

107-114.

Stoecker D.K., Taniguchi A. and Michaels A.E. (1989)

Abundance of autotrophic, mixotrophic, and

heterotrophic planktonic ciliates in shelf and slope

waters. Mar Ecol Prog Ser 50: 241-254.

Utermohl von H. (1931) Neue Wege in der quantitativen

Erfassung des Planktons. (Mit besondere

Beriicksichtigung des Ultraplanktons). Verhandlungen



der Int. Vereinigung fur Theor. und Angew. Limnol. 5:

567-595.

Wikner J. and Hagström Å. (1999) Bacterioplankton intra-

annual variability at various allochthonous loading:

Importance of hydrography and competition. Aquat.

Microb. Ecol. 20: 245-260.

Zubkov M.V., Burkill P.H. and Topping J.N.(2007). Flow

cytometric enumeration of DNA-stained oceanic

planktonic protists. J. Plankton Res. 29: 79–86.

doi:10.1093/plankt/fbl059.

Co-funded by the European Union D4.1 Standard Operating Procedures| 4 Periphyton


4. Periphyton

4.1 Definition and Terms

DW Dry Weight


The materials needed:

✓ Demineralized water

✓ Light sensor (for example: Licor: LI-250A)

✓ Microbalance (d=0.001mg)

✓ Light source (≥36 Watt)

✓ Oven (60 °C)

✓ Vacuum filtration system

✓ Desiccator

✓ GF/F-filter (ø=25mm and pore size =0.7 µm)

✓ Transparent polypropylene sheets

✓ Petri disks (ø=55mm)

✓ Scalpel

✓ Weighing paper

4.3 Health and safety regulation

Some remarks about ‘Health and safety of field operators (practical advice)’ can be found in (1):

“1. Always wear thigh waders or some other form of protection for your feet.

2. Always wear a life jacket while sampling.

3. Never sample in parts of the river which are out of your depth.

4. When sampling rivers which may be heavily polluted or polluted with fecal matter, be sure to always wear

gloves. "

4.4 Environmental Indications

A plan for the disposal of mesocosm waste needs to be prepared prior to the experiments. The plan must be


European Commission. The Safety Data Sheets (SDSs) need to be revisited for the disposal of reagents and

chemicals prior to waste disposal.





4.5 Methods

The following describes the periphyton protocol of sampling periphyton on artificial substrates as used by

the NIOO-KNAW. Here an artificial substrate is used and it is assumed that you do not need to know the exact

community composition of the periphyton. Periphyton can also be sampled from natural substrates or other

types of artificial substrates as described e.g. in

https://archive.epa.gov/water/archive/web/html/ch06main.html. However, sampling of natural substrates,

and artificial substrates other than described in this SOP, is beyond the scope of this document

4.5.1 Prior to sample collection

4.5.1.1 Required equipment

The sampling equipment and supplies shall be provided by the AQUACOSM host facility. The following is a

list of equipment and supplies that can be used for recordkeeping, and sampling

For Recordkeeping:

✓ Overview of mesocosm labels & treatments

✓ Outline of sampling design (strata to be sampled)

✓ Field sheets (Metadata Sheets) (Field protocol; DQ from [2])

✓ Permanent markers and pens/pencils

✓ Other relevant paper work

✓ Notebook for contemporaneous notes if required

The Sampling Equipment

✓ Waterproof digital camera

✓ 50 ml plastic test tubes with caps

✓ Wire

✓ Anchors/ steel bars

✓ Light sensor (for example: Licor: LI-250A)

Additional Equipment (can be excluded)

✓ Cooler with ice

✓ Portable pH/Conductivity/Temperature meter

✓ Soil probe

✓ Meter stick

https://archive.epa.gov/water/archive/web/html/ch06main.html



4.5.1.2 Calibration (if needed for any of the sampling equipment)

✓ Consult the manuals for the light sensor and -if needed- portable pH/Conductivity/Temperature

meter for calibration procedures

4.5.2 Periphyton collection with artificial substrate (2)

4.5.2.1 Preparation of the filters

• Rinse the filters with ±100 ml demineralized water to remove the waste of the production process

(carbon, phosphates). Make sure to have sufficient filters, more than the number of strips.

• Lay the filters on the weighing paper in clean and coded Petri dishes and put these into an oven of

60 degrees °C for at least 24 hours.

• Remove the filters from the oven and let them cool down for a minimum of 45 minutes in a

desiccator.

• Weigh the filters on a microbalance and write down the weighs (DWA) of the dry filters.

4.5.2.2 Construction of the strips

• Cut the sheet of polypropylene (A4 format) over the length in 2 pieces (Fehler! Verweisquelle konnte

nicht gefunden werden.)

• Cut the strips with the dimensions of 2.0 x 16.0 cm (Figure 4-2).

o Please be mindful that the dimensions of all strips should be equal, because the biomass

growth will be expressed per surface.

• Roll the sheet into a tube and secure the upper edges (Fehler! Verweisquelle konnte nicht gefunden

werden..3 red shaded parts)

• Attach a wire on three points through the upper edge, which is uncut, to make a construction that

can hang into the water (Fehler! Verweisquelle konnte nicht gefunden werden..4).

• Make a set blank strips, which will not be placed in the water. This set will be used later to measure

the background extinction. Use steps 1 and 2 (Fehler! Verweisquelle konnte nicht gefunden w

erden..2.) to make the blank strips.



Figure 4-1: stepwise approach of the construction of the periphyton strips from an A4 sheet of polypropylene. The striped lines indicate where to cut

4.5.2.3 Placing the strips

• Place the anchors/steel bars with the strips on the selected field location(s).

• Attach the strip construction to the anchors, so that the strips can move freely in the water. This is

necessary to recreate a natural situation/environment for the periphyton in which the dynamic of

the water can be important. The advice is to keep a height of 30 cm above the sediment.

4.5.2.4 Collecting the strips

• Before visiting the field location: fill tubes of 50 ml with a fixed volume of demineralized water. The

advice is to use 45 ml water because there must be space left for the strip. It is important that the

strip will be fully submersed in the water in the tube.

• Carefully raise the construction to the water surface.

• Cut the strips at the upper edge of the construction and place the strips immediately in the tubes

with demineralized water. This is necessary to prevent the dehydration of the periphyton.

• Lower the construction carefully into the water when working with repeated samples from the same

construction.

• Alternative: Use scubadiving when it is too difficult to raise the strips because of depth or excessive

plant growth. In this case, it is recommended to put the strips directly in the tubes under water.

• Take the tubes to the lab for further analysis stored at 8 °C.



4.5.2.5 Determination of the light extinction by periphyton

• Remove the strips carefully from the tube with water. Attention: the extinction of light by periphyton

must be determined on the same day as the harvest of the strips.

• Place the strip on a predetermined fixed distance from a constant light source.

• Determine the incoming light irradiation in front of the strip with a light sensor (Figure 4-2,

measurement 1)

• Determine the transmitted light behind the strip (Figure 4-2, measurement 2).

• Measure the transmitted light using a blank strip (Figure 4-2, measurement 2).

• Determine the percentual light extinction by periphyton, using the extinction of the strip corrected

by the extinction of the blank strip.

•

𝐵𝑙𝑎𝑛𝑐 𝑐𝑜𝑟𝑟𝑒𝑐𝑡𝑖𝑜𝑛 = 𝐼𝑛𝑐𝑜𝑚𝑖𝑛𝑔 𝑙𝑖𝑔ℎ𝑡 − 𝑡𝑟𝑎𝑛𝑠𝑚𝑖𝑡𝑡𝑒𝑑 𝑙𝑖𝑔ℎ𝑡 𝑜𝑓 𝑏𝑙𝑎𝑛𝑐

𝐸𝑥𝑡𝑖𝑛𝑐𝑡𝑖𝑜𝑛 𝑏𝑦 𝑝𝑒𝑟𝑖𝑝ℎ𝑦𝑡𝑜𝑛 = 100 − (𝐼𝑛𝑐𝑜𝑚𝑖𝑛𝑔 𝑙𝑖𝑔ℎ𝑡

𝑇𝑟𝑎𝑛𝑠𝑚𝑖𝑡𝑡𝑒𝑑 𝑙𝑖𝑔ℎ𝑡 + 𝑏𝑙𝑎𝑛𝑐 𝑐𝑜𝑟𝑟𝑒𝑐𝑡𝑖𝑜𝑛∗ 100)

Figure 4-2: Setup for the determination of incoming and transmitted light of a strip overgrown with periphyton.

4.5.2.6 Determination of the periphyton biomass

• Remove the strips carefully from the tube with water. Attention: The first steps for the biomass

determination should be done on the same day as the harvest of the strips.

• Clean both sides of the strip with a scalpel in the water of the 50 mL tube.



• Filtrate the water with the scraped periphyton over the filters (see Preparation of the filters) on a

vacuum filtration system. Attention: It is possible that the filter cannot handle the total volume as

the filter becomes clogged with material. To resolve this issue, repeat the filtration step with a

smaller subsample. Please remember to write down the volume of the subsample.

• Put the filters with the filtered material back in the coded petri dishes and place the dish in an oven

for at least 24 hours on 60 °C.

• Remove the filters from the oven and let them cool down for 45 minutes in a desiccator.

• Weigh the filters on a microbalance and write down the weights (DWB) of the dry filters with the

periphyton.

• Calculate the biomass dry weight of the periphyton with the following formula:

𝐷𝑊𝑝𝑒𝑟𝑖𝑓𝑦𝑡𝑜𝑛 = 𝐷𝑊𝐵 − 𝐷𝑊𝐴

• If desired, further analyzes of the filter can be done such as for chlorophyll extraction, cyanotoxin

extraction, or C, N and P determination of the periphyton.

4.6 Quality Assurance and Quality control

Quote from (3):

“QUALITY CONTROL (QC) IN THE FIELD

1. Sample labels must be accurately and thoroughly completed, including the sample identification code,

date, stream name, sampling location, and collector's name. The outside and any inside labels of the

container should contain the same information. Chain of custody and sample log forms must include the

same information as the sample container labels. Caution! Lugol's solution and iodine-based preservatives

will turn paper labels black.

2. After sampling has been completed at a given site, all brushes, suction and scraping devices that have

come in contact with the sample should be rubbed clean and rinsed thoroughly in distilled water. The

equipment should be examined again prior to use at the next sampling site, and rinsed again if necessary.

3. After sampling, review the recorded information on all labels and forms for accuracy and completeness.

4. Collect and analyze one replicate sample from 10% of the sites to evaluate precision or repeatability of

sampling technique, collection team, sample analysis, and taxonomy.”



4.7 References 4 - Periphyton

1. Taylor JC, Harding WR, Archibald CGM. A methods

manual for the collection, preparation and analysis of diatom

samples. WRC Report No TT 281/07 Water Research

Commission, Pretoria. 2007:11-23.

2. Teurlincx S, de Senerpont Domis L. Measurement of

growth and light extinction by periphyton under field

conditions. 2017.

3. Stevenson RJ, Bahls LL. Periphyton protocols.

Chapter 6 in: MT Barbour, J. Gerritsen, BD Snyder, and JB

Stribling (eds.), Rapid Bioassessment Protocols for use in

Streams and Wadeable Rivers. EPA/841-B-99-002. US

Environmental Protection Agency, Office of Water.

Washington, DC; 1999.

From the phytoplankton SOP:










pursuant to Article 1(4) of Council Directive

91/689/EEC on hazardous waste (notified under

document number C(2000) 1147) (Text with EEA

relevance) (2000/532/EC),” Off. J. Eur. Communities,

vol. 69, 2000.

Co-funded by the European Union D4.1 Standard Operating Procedures| 5 Water Chemistry


5. Water Chemistry


Alkalinity capacity of water to resist changes in pH that would make the water more acidic

Analyte The constituent or characteristic of a sample to be measured

Aphotic Zone Zone within a water body where photosynthetic production is not possible (gross

primary production < respiration)

Blank A blank contains little to no analyte of interest. It is included in measurements to

trace contaminations or signal drift.

Conductivity Inverse of electrical resistivity, increases with ions in solution and is temperature

dependent

Detritus Dead particulate organic material

Euphotic Zone Zone within a water body where photosynthetic production is possible (gross

primary production > respiration), it roughly corresponds to 2-2.5-times of the

transparency (Secchi depth) [1]

Macrophytes Water plants

Matrix Components of a sample other than the analyte

Phytoplankton Community of free-floating, predominantly photosynthetic protists and

cyanobacteria in aquatic systems, (in limnological analysis commonly excluding

ciliates). [1]

Seston Organisms and non-living matter swimming or floating in water

Standard (analytical) Standardized reference material containing a known amount of the analyte, used to

calibrate measured signal against analyte concentration

Stratification Formation of a vertical temperature gradient within a water column that due to

differences in density avoids vertical mixing

Turbidity Cloudiness or haziness of a fluid

Zooplankton Community of free-floating, heterotrophic organisms





In this section, general guidance on the protection of health and safety while sampling and analysing water

samples from mesocosm experiments will be provided to minimize the risk of health impacts, injuries and

maximize safety. The users of this SOP are expected to be familiar with the Good Laboratory Practice (GLP)

of World Health Organization (WHO) [4] and Principles on GLP of Organisation for Economic Co-operation

and Development (OECD) [5]. Health and Safety Instructions of the mesocosm facility, if there are any, shall

be followed properly to protect the people from hazardous substances and the harmful effects of them.


site or in the laboratory), the work should be practiced consistent with national and EU regulations (see the

OSH Framework Directive 89/391/EEC, [6]). Other regulations and guidelines can be found on the EU – OSHA


shall be taken by the users of this SOP and the scientist-in-charge shall ensure that those measures comply


The table below summarizes the hazards, risks and safety measures for laboratory studies on water

chemistry.


Occupations at

risk


Laboratories o Exposure (skin, eye,

inhaling) to harmful

chemicals

o Ingestion of harmful

chemicals

o Appropriate personal protection equipment

(gloves, goggles, lab coat)

o Ventilated working area

o Being informed about risks of applied

chemicals and providing SDSs in the facility

o Clear labelling of all chemical containers

o No eating or drinking in the lab

o Washing hands after leaving the lab

o No storage of sample/ chemicals in empty

food containers

o Keeping solvents away from heat sources

(ovens, open flame, autoclave)








o Flame/ Explosion

o Hot surfaces/ steam

o Vacuum/ Implosion

o Overpressure

o Accidental release of

substances harmful to

the environment

o Use of tightly closed boxes to store solvents in

refrigerators

o Using heat – insulating gloves

o Using autoclaves with safety interlock

o Using exclusively thick-walled containers

approved for vacuum applications

o Using autoclaves with safety interlock

o Collection of chemicals and samples and

disposal according to national and local

regulations

5.2.2 Safety Instructions

Personnel involved in practical work, i.e. installation, sampling, analysis at an AQUACOSM facility has to

receive a safety instruction of the respective institute. The safety instruction summarizes rules, information

and advices related to work safety based on legislation and experience. The safety instruction needs to be

completed prior to practical work.

5.2.3 Working and Personal Protection (Safety) Equipment

Personal protective equipment (gloves, safety glasses, lab coat) has to be provided in the labs and must be

used for handling chemicals.

Gloves should be chosen according to permeation time which depends on material and thickness of gloves

and type of chemical to be used. Check lists of permeation time provided by the distributor of gloves to

choose the type of glove offering best protection. Still, some chemicals easily penetrate any kind of available

glove material. In this case gloves need to be changed immediately after contact with the chemical. Consult

the SDS for recommendations on which material of glove to use.

5.2.4 Use, Storage and Disposal of Reagents and Chemicals

Before using a chemical the first time the Safety Data Sheet (SDS) needs to be consulted to be informed about

the hazard potential of the substance. For each chemical a SDS in its most recent version has to be provided

by the distributor. In addition, SDS for all chemicals used in a lab should be made available to all users, i.e.



hard copies collected in a folder. The SDS provides information on safe handling, storing and disposing of a

chemical. Based on the hazard classifications the user has to apply appropriate personal protective

equipment and needs to work under the chemical fume hood if required to eliminate the risk of exposure.

Chemicals should be stored in original containers if possible. Other containers than original, including

samples containing chemicals need to be labelled clearly. The label should indicate name and concentration

of chemicals as well as date and person responsible. All containers need to be closed tightly and stored in a

ventilated area. Hazardous chemicals have to be stored in specific cabinets separated from other chemicals

and meeting the safety requirements of the respective hazard class. This applies for flammables, explosives,

oxidizing chemicals, acids, bases, and toxic chemicals.

Only the quantities of daily consumption should be stored directly at the working area. Corrosive chemicals

must never be stored above eye height.

Collection vessels for disposal must be clearly labelled with a systematic description of their contents. To

avoid dangerous chemical reactions, consult the SDS before mixing chemicals. Entrust waste chemicals to the

appropriate authorities for disposal.

5.2.5 Use, Storage and Disposal of the Equipment

Consult your local head of the lab about rules for disposal of equipment.

5.3 Environment Indications

A plan for the disposal of chemical waste needs to be prepared prior to the experiments. The plan must be


European Commission. The SDS needs to be revisited for the disposal of reagents and chemicals prior to

waste disposal.

5.4 Sampling

5.4.1 Introduction

Good quality of analytical data relies on (1) representative sampling, (2) suitable storage conditions, and (3)

accurate and precise measurements.

“Progress in analytical protocols results in the taking of samples increasingly becoming quality-determining

step in water quality assessment. Poor sampling design or mistakes in sampling technique or sample handling

during the sampling process inevitably lead to erroneous results, which cannot be corrected afterward.”

(Handbook of water analysis, Nollet, L., M., L., De Gelder, L., S., P., 2014)





5.4.2 Sampling Strategy/ Sampling Plan

Prior to a sampling event a sampling strategy has to be defined in consideration of selected analytes and aims

of the study. This includes specification of analytes, spatial aspects (surface sample or depth-integrated

sample), sampling frequency, sampling equipment, volume and number of samples, type and size of sample

containers, processing and storage of sample, sample coding, and standardized documentation.

Figure 5-1: Elements of Sampling Strategy (from: Practical guidelines for the analysis of seawater [2])

A. Mixed mesocosms


entire mesocosm. Here, samples should be taken near the centre of the mesocosm. Care should be taken to



avoid macrophytes, if present, while sampling. If it is unclear whether the whole water volume is efficiently

mixed, an initial series of samples can be taken in transects across the mesocosm and along the vertical axis.



careful considerations on the water layer(s) to be sampled (e.g., sampling discrete water layers; sampling

multiple depths with subsequent pooling to one combined representative sample; utilization of tube

sampler). The absence of vertical temperature profiles indicates vertical mixing, but NOT necessarily

homogenous distribution of motile organisms like (micro-) zooplankton. Hence, especially particulate matter

(chlorophyll-a, particulate nutrients) may be non-homogenously distributed even in the absence of a

thermocline, and require careful consideration of the appropriate sampling design.

5.4.3 Equipment for Sampling

The following specific pieces of equipment are suggested for collecting water samples from mesocosms

✓ Appropriate water sampler, depending on the type of sampling (stratified or integrated), and (e.g.

Schroder/Schindler/Ruttner sampler for single strata; tube sampler for depth integrated samples)

✓ As an alternative to a water sampler a sample can be retrieved by pumping, i.e. by a silicone tube

connected to a carboy which in turn is connected to a vacuum pump. Land based mesocosms can be

equipped with sampling ports, avoiding the risk of contamination from sampling equipment.

✓ Clean & rinsed sampling containers in the field

✓ Sampling containers needs to be labelled properly in the laboratory prior to sampling. The labels on

the sampling bottles need to be standardized and provide information on name of the experiment,

sampling date, mesocosm ID, and possibly depth with appropriate abbreviations of the treatments

of the experiments. Labels can be either printed or written using a permanent waterproof marker

Figure 5-2: Devices for water sampling - Ruttner Sampler, Schindler Patalas, and Tube Sampler



5.4.4 Techniques for representative sampling

Representative sampling – samples need to be representative for the mesocosm unit of interest. In case

multiple parameters (e.g. dissolved & particulate nutrients, chlorophyll-a) are sampled on the same event, it

is mandatory that all parameters are analysed from the same water sample (multiple sub-samples from one

water sample). This implies that homogenous distribution of particles (phytoplankton, bacteria, detritus) is

ensured whenever a sub-sample is taken from the sampling container.

A. Mixed mesocosms


entire mesocosm. Here, samples should be taken near the centre of the mesocosm as stated in both [10] and

[12]. Care should be taken to avoid macrophytes, if present, while sampling. If it is unclear whether the whole

water volume is efficiently mixed an initial series of samples can be taken in transects across the mesocosm

and along the vertical axis (see point 6.4.B below).



careful considerations on the water layer(s) to be sampled (e.g. sampling discrete water layers; sampling

multiple depths with subsequent pooling to one combined representative sample; utilization of tube

sampler). Absence of vertical temperature gradients indicates vertical mixing, but NOT necessarily

homogenous distribution of motile organisms like (micro-) zooplankton. Heterogeneous distribution of

phytoplankton must also be assumed if mesocosms contain structuring elements (such as macrophytes), and

has not been proven to be (practically) homogenous (see 6.4.A). C. Composite and discrete sampling

Careful consideration of the sampling design is required in case the mesocosms are stratified and include an

aphotic zone (mesocosm depth > Zeu3). In this case the integrated water sample would typically be taken from

the euphotic zone (Zeu).

For shallow mesocosms containing sediment and possibly macrophytes, a detailed sampling design for

collecting samples at multiple horizontal and vertical positions might be needed [12]:

Best Practice Advice: “The entire water column, from the surface to approx. 5 cm above the sediment, is

sampled from three positions in each enclosure: 10, 30 and 60 cm from the enclosure wall. Two samples are

prepared: one to be used for chemical and phytoplankton analyses where water is sampled without touching

the plants – and one to be used for zooplankton analysis, where water is sampled also close to the plants.”

“The best way to sample from the entire water column is by using tube samplers which sample from top to

bottom. The diameter of the tube should not be too small to avoid zooplankton escaping during sampling (>

6 cm). If it is not possible to use a tube sampler, samples can be taken with a Ruttner water sampler from the

surface (20 cm below the water surface), middle and the bottom (20 cm above the sediment). The sample in

the middle should be adjusted according to the enclosure type and actual water depth [11].

3 Zeu corresponds to 2-2.5 x Secchi depth



5.4.5 Quality Assurance Considerations

Any treatment of a sample, like transfer into another container, preservation, filtration, dilution may

introduce a contamination or alter the sample in another way. Consequently, the ideal procedure would be

to transfer the sample directly into the container for storage and analyse it immediately.

Sampling Technique

Already the technique of sampling can alter the quality of a sample. Pumping or temperature change may

already affect concentration of dissolved gases.

Sampling Sequence

Following sequence is recommended for subsampling water from a water sampler:

O2, pH/DIC/Alkalinity/ Nutrients

Contaminations

Potential sources for contamination of samples throughout the entire sampling procedure need to be

identified and avoided. Problematic contaminations can be either the chemical compound of interest or any

other substance interfering with the chemical analysis.

Sources for contaminations may be:

• Water Sampler/ Tube: Make sure it is clean and the materials suite your purpose.

• Sample Vials: Materials may release/ adsorb compounds into/ from your sample. Use clean

containers of appropriate material and consider rinsing them with sample first.

• Skin: Wear gloves to avoid contaminations with sweat, remains of soap, sun screen, etc.

• Boat Exhaust/ Cigarette Smoke: Rich in ammonia

Best Practice Advice: Use silicone tubing for O2, pH, DIC

Sample Matrix

The matrix of a sample (turbidity, salinity, colour, alkalinity, other chemical constituents) can affect the

chemical analysis in various ways. Optical characteristics of the sample can influence measurements based

on absorbance and fluorescence or may interfere with the signal-producing reaction. If sample and standard

do not share the same matrix, the calibration is incorrect.

Turbidity: If possible, turbidity should be removed by filtration. Alternatively, the chemically untreated

sample can be used as blank to correct for turbidity.

Salinity: Sea salt may suppress analyte absorbance in spectrophotometric measurements like phosphate,

silicate and ammonia. This problem can be addressed by preparing reference solutions of standards with



nutrient-free or low-nutrient sea water, or artificial sea water. The salinity should be equal to that of the

samples. If salinity varies a lot, a mathematical correction of salt effects can be applied by establishing a

correction function for each analyte showing salt effects.

5.5 Filtration

5.5.1 Introduction

By means of filtration, the water sample is separated into a particulate and dissolved fraction for separate

analysis. The water sample can be forced through the filter material either with vacuum or over-pressure.

The filter type affects minimal particle size retained and filtering capacity. Filter material needs to be chosen

to minimize interaction with the analyte and to allow required cleaning procedures (see 7.5 and 7.6). Any

meaningful analysis of a set of filtered samples must include at least 4 blanks. Blank filters should be cleaned

according to established procedures and from the same batch as the filters used for the samples.

5.5.2 Vacuum Filtration

The circulate filter is placed between the filter support and the funnel held in place with a clamp. Filtrate is

collected in the receiving flask while particles are retained on the filter. Applied vacuum should not exceed

200 m bar to avoid rupture of cells and leaching of particulate material [13].

Figure 5-3: Vacuum filtration (from: Practical guidelines for the analysis of seawater [2])



5.5.3 Pressure Filtration

As an alternative to vacuum filtration a sample can be pressurized and passed through a filter. Filtration with

syringe and filter holder may be an optimal procedure if samples need to be filtered immediately. Required

equipment is small and filtration does not depend on infrastructure, hence filtration with a syringe can be

easily performed in the field. Still it is only appropriate when rather small volumes of sample need to be

filtered.

To minimize sample carry-over syringe and filter need to be flushed with new sample, especially because

filter holders contain some dead volume.

Figure 5-4: Disposable filter discs and reusable filter holders used for filtration with the syringe

5.5.4 Filter Types

Glass fibre filters are best choice for organic carbon (DOC/POC), nitrogen (DON/PON), and phosphorus

(DOP/POP). Glass fibre filters have a poor uniform pore size, but they can be easily cleaned by baking at high

temperature (typically 450°C) for several hours to produce low blanks for these elements and at the same

time they provide good flow rates for high-volume samples. Glass fibre filters are the classical filter material

for the determination of chlorophyll pigments and are also suitable for the filtration of nutrient samples

except silica, in which polycarbonate filters are mostly used [2].

Besides glass fibre filters (GF/F ~ 0,7µm and GF/C ~ 1,2µm) membrane filters of 0,45 and 0,2µm pore size are

used to separate particulate and dissolved phases of water samples [3].




● For efficient retention of seston including cyanobacteria GF/F filters should be used.

● Filters used for quantification of particulate carbon should be pre-combusted at 450°C

● Filters for quantification of particulate phosphorus should be acid washed.

Table 5-2: Filter Materials and their characteristics (from: Practical guidelines for the analysis of seawater [2])

5.5.5 Cleaning procedure for glass fibre filters

To assure glass fibre filters are free of organic traces they have to undergo a cleaning procedure.

Bake filters at 450°C for 4 hours. Place filters in a glass beaker and cover them with diluted acid (i.e. 10-15%

Hydrochloric Acid). Rinse filters repeatedly with analytical grade water and dry them at 60°C.



✓ Best practice advice: Apply the above cleaning procedure (acid washing/ baking) to all glass fibre

filters even if not required for the subsequent analysis (i.e. chlorophyll). This will assure identical

filtration results/ comparability of chemical parameters since the high temperature and washing can

affect the effective pore size of the filters.

5.5.6 Cleaning procedure for synthetic membrane filters

Rinse with analytical grade water (i.e. MilliQ) and allow the filters to stand soaked in MilliQ for 30 minutes,

rinse again with MilliQ and press out the remaining water with air. A test tube rack can be used as support.

Always discard the first millilitres of sample to waste.

✓ Best practice advice: Use syringes with plastic plungers, avoid syringes with rubber plungers. The

rubber or the grease on the rubber is a potential source for contamination.

5.5.7 Volume for filtration

The water volume required in order to collect sufficient material on a filter depends both on the parameter,

the sensitivity of the methodology, and esp. on the density of particles in the water, which in turn depends

on the trophic state of the experimental system.

✓ As a rule of thumb, a clearly visible coloration on the filter (well visible against the white background

of a glass fibre filter) ensures reasonable quantity of material for most common analytical protocols

(PON/C/P, Chlorophyll-a).

✓ A careful documentation of filtered volume is mandatory in order to calculate concentrations (e.g.

µg Chl-a L-1).

5.6 Sample Storage

The biological activity in water does not stop with sample collection, since bacteria and micro- and nano-

plankton continue to digest and excrete material [13].

Nutrients are subject to rapid changes in their concentration within a few hours in unpreserved samples [2].

Sample preservation is needed whenever measurements cannot be performed immediately or when a

backup for potential reanalysis is required.

Each analyte has its own reaction chemistry and consequently different requirements for storage in solution.

Therefore, no general procedure can be recommended for the storage of water samples [3].

Optimum storage conditions differ largely among parameters (see Table 5-3 below).



Table 5-3: Best Practice Advice for Storage of Water Samples

Analyte Container Preservation Notes

Soluble Reactive

Phosphorus

Glass, acid washed

LDPE, acid washed

Filter and measure immediately or store ≤4°C in the dark if

analysed within 24hrs.

For longer storage freeze filtered samples at -20°C.

Calcareous water (Ca2+ > 100mgL-1)

must be acidified with 1ml

concentrated hydrochloric acid L-1

before freezing to prevent co-

precipitation of phosphate

Particulate Phosphorus Plastic petri dish or

Eppendorf tube

Filter immediately, freeze filters at -20°C

Total Phosphorus Glass, acid washed

LDPE, acid washed

Store in containers and volumes desired for digestion ≤4°C in

the dark.

For long term storage freeze samples at -20°C.






Total Dissolved

Phosphorus

Glass, acid washed

Store filtered samples in containers and volumes desired for

digestion ≤4°C in the dark.

For long term storage freeze filtered samples at -20°C.






Total Reactive

Phosphorus

Glass, acid washed

Measure immediately, or store at 4°C in the dark if analysed

within 24hrs.

Ammonia Plastic or Glass Measure immediately, if necessary filter and store at 4°C for

up to 24hrs, or filter and freeze unacidified at -20°C for up to

28d

Note that samples that have been

measured immediately are not

necessarily comparable with

samples that have been filtered and

frozen. this is especially true for

ammonia.

DOC (NPOC) Polycarbonate or

cell culture flasks

Filter through combusted GF/F or 0,2µm membrane filter into

combusted glass vial and preserve immediately with H3PO4,

store in the dark at 4o C.

Nitrate Plastic or Glass Measure immediately, if necessary filter and store at 4°C up to

2d. For longer storage filter and store at -20°C.

In samples preserved with acid, NO3-

and NO2- cannot be determined as

individual species.

Nitrite Plastic or Glass Measure immediately, or filter and store at -20°C for longer

storage

Never acidify for storage.

Total Nitrogen Plastic or Glass Acidified to pH 1-2 with H2SO4 samples can be stored 1 month

Chlorophyll a Plastic petri dish or

Eppendorf tube

Filter immediately after sampling and freeze at -20C Protect from light, ideally store at -

80°C

Dissolved Inorganic

Silicate

Plastic Filter immediately. Store at 4°C for up to 1 month. Do not acidify, silicate precipitates

under acidic conditions. Avoid

freezing.

Total alkalinity Glass, acid washed Filter immediately after sampling and store at 4°C up to 24h or

up to a month if the samples is poisoned.

Samples can be poisoned with HgCl2

solution.

Oxygen, dissolved Plastic or Glass Cool, protect from air and light, store up to 6hrs On-site measurement preferable



pH Plastic or Glass Cool, protect from air (fill bottle to cap) On-site measurement preferable

Alkalinity Plastic or Glass Cool, protect from air (fill bottle to cap), store up to 24h On-site measurement preferable,

especially for samples high in

dissolved gases

Best practice advice: Samples preserved by acidification should be neutralized prior to analysis. Acid and

base may introduce background to the sample, hence standards used in subsequent measurements must

undergo the same treatment.

5.7 Auxiliary measurements

Here we briefly outline measurements that are often conducted in combination with sampling for water

chemistry. Transparency and basic physical parameters are often taken alongside water sampling for water

chemistry and phytoplankton.

Best practice advice: The relevance of measuring irradiation and transparency depends on the depth of the

mesocosms. In shallow mesocosms (d < 2m), light very likely will not be limiting during the growing season.

Moreover, the influence of the suspended particles including phytoplankton on under water light climate is

very limited in shallow water columns.

5.7.1 Water transparency (aka Secchi depth)

Water transparency is a key parameter in limnology and oceanography, informing especially about the optical

depth of the water column, which affects e.g. the vertical structure of biota and biological processes, such as

primary production, but also vertical migration of the zooplankton.

Transparency is typically measured by lowering a white disk vertically into the water until it is not visible

anymore. Now the disk is slowly lifted, until it becomes visible. The depth where the disk just becomes visible

is defined Secchi depth. A detailed outline is given in

http://www.helcom.fi/Lists/Publications/Guidelines%20for%20measuring%20Secchi%20depth.pdf.

5.7.2 Light measurements

Light is a key resource for primary production. In aquatic ecology, it is commonly measured as photo-

synthetically active radiation (PAR; https://www.licor.com/documents/liuswfuvtqn7e9loxaut ). For proper

quantification of light, irradiation needs to be measured by an appropriate probe at various positions inside

the mesocosm, taking the optical structure (illuminated vs. shaded side etc.) and day-time into account.

Especially in narrow mesocosms with opaque walls, a proper quantification of irradiation may be very

difficult.

https://www.licor.com/documents/liuswfuvtqn7e9loxaut



If light intensity is a key parameter and e.g. manipulated by shading, comparing irradiance at a fixed position

(e.g. middle of water column) may be a good proxy. An example for light-manipulation inside mesocosms

can be found here https://www.nature.com/articles/srep29286

5.7.3 Standard physical parameters (Temperature, Oxygen concentration, Conductivity)

Standard physical parameters, especially water temperature, conductivity, pH and oxygen concentration are

often measured using handheld probes. For measuring all of these parameters, the same recommendations

regarding representativeness apply, as outlined above (→ 5.4.4 Techniques for representative sampling). If

mesocosms are stratified, layers must be sampled separately. Measurements of physical parameters using

submersible probes are described in section 6.4 Field Parameters and Sensors (High Frequency

Measurements).



5.8 Methods applied by Project Partners

5.8.1 Aarhus University (AU)

Parameter

Detection Mode

Method Name Reference Comments Manual Automated

Alkalinity X Titration DS 253, 1977

Chlorophyll a X

Spectrophotometric determination in ethanol

extract

DS 2201, 1986

X Chlorophyll a sensors - Turner designs - Cyclops 7F

Conductivity X YSI 6600 Xylem Analytics

Nitrogen

NH4

X

Photometric method, ammonia-nitrogen

(indophenol blue)

DS 224, 1975

NO3

X

Automated Hydrazine Reduction method with FIA

Star 5000 Foss FIA Star 5000

TN

X

Automated Hydrazine Reduction method with FIA

Star 5000, digestion with peroxodisulfate/NaOH

DS 221/Foss FIA Star 5000

Oxygen X Oxygen probe Oxyguard

pH X pH probe Oxyguard

Phosphorus



SRP

X

Orthophosphate photometric method (ascorbic

acid/molybdate) DS 291, 1985

TP

X

Total Phosphorus photometric method (ascorbic

acid/molybdate), digestion with peroxodisulfate DS 292, 1985

Silica X Photometric method, molybdosilicate DMU T.A. nr. 22

5.8.2 Centro de Biodiversidade e Recursos Genéticos – Universidade de Évora (CIBIO)

Parameter

Detection Mode Method Name

Reference

Comments

Manual Automated

Conductivity X Digital rugged conductivity probe CDC40105, Hach

Oxygen X

Digital, luminescent/optical dissolved

oxygen (LDO) probe LDO101, Hach

pH X

Digital combination pH electrode with

built-in temperature sensor PHC101, Hach

Chlorophyll a X Handheld Fluorometer /Chlorophyll in vivo Aquafluor®, Arar 1997 (EPA Method 445.0)

No cell disruption and

acidification is applied

Nitrogen

NH3 X Indophenol Blue Method Ivancic & Deggobis 1984

NO2 X Automated Hydrazine Reduction Method ISO 13395:1996




TN X

Persulfate Digestion, Hydrazine Reduction

Method

Clesceri 1999 (4500-P, chapter J);

ISO 13395:1996

Phosphorus

TP/ TDP X

Persulfate Digestion and Ascorbic Acid

Method Grasshoff 1999 (chapter 10.2.13)

References

Arar, E. J., & Collins, G. B. (1997). Method 445.0: In vitro determination of chlorophyll a and pheophytin a in marine and freshwater algae by fluorescence.

Cincinnati: United States Environmental Protection Agency, Office of Research and Development, National Exposure Research Laboratory.

Clesceri, L. S., Greenberg, A. E., & Eaton, A.D. (1996). Standard methods for the examination of water and wastewater. APHA, AWWA and WPCF, Washington DC.

DIN EN ISO 15681-2, Water quality - Determination of orthophosphate and total phosphorus contents by flow analysis (FIA and CFA) - Part 2: Method by

continuous flow analysis (CFA), 06-2001



5.8.3 MARine Biodiversity, Conservation and Exploitation (CNRS-MARBEC)

Parameter

Detection Mode


Chlorophyll a X Fluorometric Detection Strickland and Parson 1972

Acetone extraction

Ultrasonic cell disruption

Pigments X High-Performance Liquid Chromatography Method Zapata et al. 2000

Suspended

particulate matter

(SPM)

X Gravimetric Strickland and Parson 1972

Nitrogen

NH4 X Ortho-phthaldialdehyde fluorometric Method (OPA) Holmes et al. 1999

NO2

X

CFA-based photometric detection, diazotization of

NO2 (Gries-Ilosvay reaction) with sulphanilamide

produce a reddish-purple colour, which is measured

at 540 nm.

ISO 13395:1996

NO3

X

CFA-based photometric detection, nitrate reduction

to nitrite by use of coppered Cd-granules and

detection as nitrite (see above)

ISO 13395:1996

Oxygen (dissolved) X Titration – Winkler method Strickland and Parsons 1972

- use of a silicone tube for sampling

to avoid air bubbles

- use of calibrated bottles/flasks.



pHT (total scale) X

Spectrophotometric method (based on the

absorption ratio of the sulfonephthalein dye, m-

cresole purple

Byrne 1993

Liu et al 2011

- use of a silicone tube for sampling

to avoid air bubbles

Phosphorus

SRP

X

CFA-based photometric detection, reduction to

molybdenum blue complex by use of ascorbic acid.

The complex is measured at 660 nm.

ISO 15681-2

Salinity X

Silica X


molybdenum blue complex by use of ascorbic acid ISO-16264

Oxalic acid is added to avoid

phosphate interference

Temperature X

References

Byrne R.H. (1993). Spectrophotometric seawater pH measurements: total hydrogen ion concentration scale calibration of m-cresol purple and at-sea results.

Deep-Sea Research, Vol. 40, No 10, pp 2215-2129.

Holmes, R. M., Aminot, A., Kérouel, R., Hooker, B. A., & and Peterson, B. J. (1999). A simple and precise method for measuring ammonium in marine and freshwater

ecosystems. Can. J. Fish. Aquat. Sci., 56, 1801–1808

ISO 13395:1996, Water quality -- Determination of nitrite nitrogen and nitrate nitrogen and the sum of both by flow analysis (FIA and CFA) and spectrometric

detection.

ISO 15681-2, Determination of orthophosphate and total phosphorus contents by flow analysis (FIA and CFA), Part 2: Method by continuous flow analysis (CFA).

ISO-16264: Determination of soluble silicates by flow analysis (FIA and CFA) and photometric detection.



Liu X., Patsavas M.C and Byrne R.H. (2011). Purification and characterization of meta-cresol purple for spectrophotometric seawater pH measurements. Environ.

Sci. Technol., 2011, 45 (11), pp 4862–4868.

Strickland, J.D.H., & Parsons, T.R. (1972). A Practical Handbook of Seawater Analysis. 2nd Edition, Fisheries Research Board of Canada Bulletin, 167, 310 p.

Zapata, M., Rodriguez, F., & Garrido, J. (2000). Separation of chlorophylls and carotenoids from marine phytoplankton: A new HPLC method using a reversed

phase C-8 column and pyridine-containing mobile phases. Mar. Ecol. Prog. Ser., 195, 29–45, doi:10.3354/meps195029

5.8.4 Ecole Normale Superieure (ENS)

Parameter

Detection Mode


Alkalinity,

total

X Potentiometric automated titration with open cell

method

Dickson et al. 2007

Chlorophyll a X Fluorometric Detection after Acetone Extraction Jespersen and Christoffersen 1987

Dissolved oxygen X Optical method (optode sensor)

Nitrogen

NO2

X Colorimetric with Sulfanilamide and NEDD Grasshoff et al. 1983 N° G-173-96 Rev. 10 Seal analytical AA3

autoanalyzer method

X Colorimetric Strickland and Parsons, 1972 UV VIS

spectrophotometer



NO3 X

Colorimetric with Sodium salicylate Strickland and Parsons, 1972 UV VIS

spectrophotometer

X Cd reduction and colorimetric method with

Sulfanilamide and NEDD

Grasshoff et al. 1983 N° G-392-08 Rev. 5 Seal analytical AA3

autoanalyzer method

NH4+ X Ortho-phthaldialdehyde (OPA) fluorometric Method Kerouel and Amniot 1997 NH4+

TN

X Persulfate digestion, Cd reduction and colorimetric

method

Grasshoff et al. 1983

N° G-392-08 Rev. 5

Seal analytical AA3

autoanalyzer method

pH X

Potentiometric with glass/reference electrode cell

(total scale)

Dickson et al. 2007

Phosphorus

SRP X Colorimetric with molybdate and ascorbic acid Strickland and Parsons, 1972 UV VIS

spectrophotometer

X Colorimetric with molybdate and ascorbic acid Murphy and Riley 1962

Drummon and Maher 1995

N° G-175-96 Rev. 15 (Multitest MT 18)

TP/TDP

X Persulfate and sulfuric acid digestion then molybdate

and ascorbic acid method

Grasshoff et al. 1983 N° G-393-08 Rev. 4 Seal analytical AA3

autoanalyzer method

Silicate X Ascorbic Acid - Molybdenum – Oxalic - Blue Complex Grasshoff et al. 1983 N°. G-177-96 Rev. 11

(Multitest MT19)

Seal analytical AA3

autoanalyzer method



References

Dickson A.G., Sabine, C.L. and Christian, J.R. (Eds.) 2007. Guide to best practices for ocean CO2 measurements. PICES Special Publication 3, 191 pp.

Jespersen, A-M. & K. Christophersen, 1987. Measurements of chlorophyll- a from phytoplankton using ethyl alcohol as extraction solvent. Arch.Hydrobiol. 109:

445-454.

Kerouel, R.and Amniot, A. Marine Chemistry Vol. 57, no 3-4, pp.265-275, Jul 1997.

Strickland, J. D. H., and Parsons, T. R. (1972). A practical handbook of seawater analysis. B. Fish. Res. Board Can. 167, 311.

K. Grasshoff et al., Methods of Seawater Analysis, 2nd edition, Verlag Chemie, 1983.

Murphy, J. and Riley J.P., 1962. A modified single solution method for the determination of phosphate in natural waters. Analytica Chimica Acta 27:31-36.

Drummon, L. and Maher, W., 1995. Re-examination of the optimum conditions for the analysis of phosphate. Analytica Chimica Acta 302: 69-74.

5.8.5 GEOMAR Helmholtz Centre for Ocean Research Kiel (GEOMAR)

Parameter


Reference

Comments

Manual Automated

Alkalinity,

total X Potentiometric titration, open-cell method Dickson et. al., 2003

Carbon

DIC X Acidification, gas stripping, Infrared absorption e.g. Goyet & Snover, 1993



POC X

Treated with fuming HCL in a desiccator for 2h,

elemental analysis accomplished by combustion

analysis

Sharp 1974, Hansen and Koroleff 1999;

Grasshoff “Methods of seawater

analysis”, 1999

TPC X

Elemental analysis accomplished by combustion

analysis



analysis”, 2001

Chlorophyll a X Fluorometric Detection after acetone extraction Welschmeyer 1994

Pigments X

Reverse-phase high-performance liquid

chromatography Barlow et al., 1994

Nitrogen

DON X Determination by alkaline persulphate oxidation Hansen and Koroleff, 1999

NH4 X Determined fluorometrically Holmes et. al. 1999

NO2 X

Automated Camium Reduction Method,

photometrically

Murphey and Riley et.al., 1962; Hansen

and Koroleff, 1999; Grasshoff “Methods of

seawater analysis”, 1999; NIOZ –

Nederlands Instituut for Onderzoek der

Zee (Royal Netherlands Institute ), Den

Hoorn (Texel), The Netherlands

NO3 X

Automated Cadmium Reduction Method,

photometrically


and Koroleff, 1999, modified by Keroul

and Aminot 1997, Grasshoff “Methods of





Zee (Royal Netherlands Institute for Sea

Reserach), Den Hoorn (Texel), The

Netherlands

PON


analysis



analysis”, 2000

TPN

X


analysis



analysis”, 2002

pH T (total

scale) X

Spectrophotometric method (based on the

absorption ratio of the sulfonephthalein dey, m-

cresole purple Clayton and Byrne, 1993

Phosphorus

DOP X Determination by alkaline persulphate oxidation Hansen and Koroleff, 1999

SRP X Automated Cadmium Reduction Method










Netherlands

TPP X Spectrophotometrically

Hansen and Koroleff, 1999; Holmes et al.,

1999

Silica

Biogenic

silica X

Spectrophotometrically, leaching method (135

minutes, 85°C with 0.1M NaOH) Hansen and Koroleff, 1999

Silic acid X Automated Cadmium Reduction Method








Netherlands



Hellenic Center for Marine Research (HCMR)

Parameter


Reference

Comments

Manual Automated

Carbon X

POC X

CHN analyzer Hedges and Stern (1984)

TOC

X

high-temperature catalytic oxidation

method Sempéré et al. (2002)

Chlorophyll a X

Fluorometric Detection after acetone

extraction Holm-Hansen et al. (1965)

Nitrogen

NH3

X

Vis/UV spectrophotometric

determination Ivancic & Deggobis 1984

NO2

X


determination Strickland and Parsons (1972)

NO3

X



PON X

CHN analyzer Hedges and Stern (1984)

TN

X

Wet-oxidation

Pujo-Pay & Raimbault (1994) and

Raimbault et al. (1999)

Oxygen,

dissolved X Winkler Carpenter (1965a, b)



Phosphorus

SRP

X


determination Strickland and Parsons (1972) micromolar level

SRP X

MAGIC method Rimmelin and Moutin, 2005 nanomolar level

TP

X

Wet-oxidation

Pujo-Pay & Raimbault (1994) and

Raimbault et al. (1999)

Silica X



References

Carpenter, J. H., 1965(a). The accuracy of the Winkler method for the dissolved oxygen analysis. Limnology and Oceanography, 10, 135-140.

Carpenter, J. H., 1965(b). The Chesapeake Bay Institute technique for dissolved oxygen method. Limnology and Oceanography, 10, 141-143.

Hedges, J. I., and Stern, J. H. (1984). Carbon and Nitrogen determination of carbonate-containing solids. Limnol. Oceanogr. 29, 657–663.

Holm-Hansen, O., Lorenzen, C. J., Holmes, R. W., and Strickland, J. D. H. (1965). Fluorometric determination of chlorophyll. J. Cons. Perm. Int. Explor. Mer. 30, 3–

15.

Invancic, I., and Degobbis, D. (1984). An optimal manual procedure for ammonia analysis in natural waters by the indophenol blue method. Water Res. 18, 1143–

1147.

Kirchmann, D. L., Newell, S. Y., and Hodson, R.,E. (1986). Incorporation versus biosynthesis of leucine: implications for measuring rates of protein syntheis and

biomass production by bacteria in marine systems. Mar. Ecol. Prog. Ser. 32, 47–59.



Lin, P., Chen, M., and Guo, L. (2012). Speciation and transformation of phosphorus and its mixing behavior in the Bay of St. Louis estuary in the northern Gulf of

Mexico. Geochim. Cosmochim. Acta 87, 283–298.

Miyazaki, Y., Kawamura, K., Jung, J., Furutani, H., and Uematsu, M. (2011). Latitudinal distributions of organic nitrogen and organic carbon in marine aerosols

over the western North Pacific. Atmos. Chem. Phys. 11, 3037–3049. doi:10.5194/acp-11-3037-2011.

Pujo-Pay, M., Raimbault, P., 1994. Improvement of the wet-oxidation procedure for simultaneous determination of particulate organic nitrogen and phosphorus

collected on filters. Mar. Ecol. Prog. Ser. 105, 203-207.

Raimbault, P., Pouvesle, W., Diaz, F., Garcia, N., Sempere R., 1999. Wet oxidation and automated colorimetry for simultaneous determination of organic carbon,

nitrogen and phosphorus dissolved in seawater. Marine Chemistry, 66, 161-169.

Sempéré, R., Panagiotopoulos, C., Lafont, R., Marroni, B., and Van Wambeke, F. (2002). Total organic carbon dynamics in the Aegean Sea. J. Mar. Syst. 33–34,

355–364.

Smith, D. C., and Azam, F. (1992). A simple, economical method for measuring bacterial protein synthesis rates in seawater using 3H-leucine. Mar. Microb. Food

Webs 6, 107–114.

Steeman-Nielsen, E. (1952). The use of radio-active carbon (C14) for measuring organic production in the sea. Journal du Cons 18, 117–140.

Strickland, J. D. H., and Parsons, T. R. (1972). A practical handbook of seawater analysis. B. Fish. Res. Board Can. 167, 311.

Rimmelin, P., and Moutin, T. (2005). Re-examination of the MAGIC method to determine low orthophosphate concentration in seawater. Anal. Chim. Acta 548,

174–182.



5.8.6 Ludwig-Maximilians-Universität Munich (LMU)

Parameter

Detection Mode


Alkalinity X Acidic titration DIN ISO 9963-1/2

Carbon (POC) X Elemental analyser DIN 38409-46; Hedges & Stern 1984

Chlorophyll a X

In vitro: fluorometric detection after acetone

extraction DIN 38412-16

X

In vitro: fluorometric detection after ethanol

extraction DIN 38412-16

X

In vivo: fluorometric detection (Algal lab

analyser, Turner, AquaPen)

Chlorophyll content is excited

by coloured LEDs and allocated

to the different algal classes

Cl- X Ion chromatography DIN ISO 10304-1; Smith & Chang 1983

Nitrogen

NO2 X Ion chromatography DIN ISO 10304-1; Smith & Chang 1983

NO3 X Ion chromatography DIN ISO 10304-1; Smith & Chang 1983

NH4 X Fluorometrical Holmes et al. 1999

PON X Elemental analyser Hedges & Stern 1984

Oxygen, dissolved X Winkler Carpenter (1965a, b)

Phosphorus

PP X Photometric with ammoniummolybdate DIN ISO 6878:2004; Grasshoff et al. 1999



SRP X Photometric with ammoniummolybdate DIN ISO 6878:2004; Grasshoff et al. 1999

TP X Photometric with ammoniummolybdate DIN ISO 6878:2004; Grasshoff et al. 1999

Salinity X 2 graphite electrodes WTW multi probe

SiO2 X Photometric detection DIN ISO 15923-1

SO4 Ion chromatography DIN ISO 10304-1; Smith & Chang 1983

References

Carpenter, J. H., 1965(a). The accuracy of the Winkler method for the dissolved oxygen analysis. Limnology and Oceanography, 10, 135-140.

Carpenter, J. H., 1965(b). The Chesapeake Bay Institute technique for dissolved oxygen method. Limnology and Oceanography, 10, 141-143.

Grasshoff, K., Kremling, K., & Ehrhardt, M. (Eds.). (1999). Methods of seawater analysis. John Wiley & Sons

Hedges, J. I., & Stern, J. H. (1984). Carbon and nitrogen determinations of carbonate‐containing solids. Limnology and oceanography, 29(3), 657-663.

Holmes, R.M., Aminot, A., Kérouel, R., Hooker, B.A & Peterson, B.J. (1999) A simple and precise method for measuring ammonium in marine and freshwater

ecosystems. Can.J.Fish.Aquat.Sci. 56: 1801-1808.

Smith, F. C., & Chang, R. C. C. (1983). The practice of ion chromatography. Wiley.

DIN EN ISO 9963-1:1996-02: Wasserbeschaffenheit - Bestimmung der Alkalinität - Teil 1: Bestimmung der gesamten und der zusammengesetzten Alkalinität

(ISO 9963-1:1994); Deutsche Fassung EN ISO 9963-1:1995

DIN EN ISO 9963-2:1996-02: Wasserbeschaffenheit - Bestimmung der Alkalinität - Teil 2: Bestimmung der Carbonatalkalinität (ISO 9963-2:1994); Deutsche

Fassung EN ISO 9963-2:1995

DIN 38409-46:2012-12: Deutsche Einheitsverfahren zur Wasser-, Abwasser- und Schlammuntersuchung - Summarische Wirkungs- und Stoffkenngrößen

(Gruppe H) - Teil 46: Bestimmung des ausblasbaren organischen Kohlenstoffs (POC) (H 46)



DIN 38412-16:1985-12: Deutsche Einheitsverfahren zur Wasser-, Abwasser- und Schlammuntersuchung; Testverfahren mit Wasserorganismen (Gruppe L);

Bestimmung des Chlorophyll-a-Gehaltes von Oberflächenwasser (L 16)

DIN EN ISO 10304-1:2009-07: Wasserbeschaffenheit - Bestimmung von gelösten Anionen mittels Flüssigkeits-Ionenchromatographie - Teil 1: Bestimmung von

Bromid, Chlorid, Fluorid, Nitrat, Nitrit, Phosphat und Sulfat (ISO 10304-1:2007); Deutsche Fassung EN ISO 10304-1:2009

DIN EN ISO 6878:2004-09: Wasserbeschaffenheit - Bestimmung von Phosphor - Photometrisches Verfahren mittels Ammoniummolybdat (ISO 6878:2004);

Deutsche Fassung EN ISO 6878:2004

DIN ISO 15923-1:2014-07: Wasserbeschaffenheit - Bestimmung von ausgewählten Parametern mittels Einzelanalysensystemen - Teil 1: Ammonium, Nitrat, Nitrit,

Chlorid, Orthophosphat, Sulfat und Silikat durch photometrische Detektion (ISO 15923-1:2013)

5.8.7 Middle East Technical University (METU)

Parameter


Reference

Comments

Manual Automated

Alkalinity X Titration

Standard Methods, 22. Edition. American

Health Association, 1996.

Chlorophyll a X

Spectrophotometric Detection after dissolving in

ethanol

Jespersen, A-M. & K. Christophersen,

1987.

Nitrogen

NH3 X The Skalar Autoanalyzer method

San++ Automated Wet Chemistry

Analyzer, Skalar Analytical,



B.V., Breda, The Netherlands

X Indophenol Blue Method Chaney, A. L. and Morbach, E. P., 1982.

NO2 X The Skalar Autoanalyzer method




X Spectrophotometric method (pink dye)

Mackereth, F.J., H. J. Heron & J. F. Talling,

1978.

NO3 X The Skalar Autoanalyzer method




X Spectrophotometric method (pink dye)


1978.

TN

X The Skalar Autoanalyzer method




Phosphorus

SRP X Ascorbic Acid Method


1978.

TP/ TDP X Persulfate Digestion, Ascorbic Acid Method


1978.

Silica X

Molybdosilicic Acid Method/ Heteropoly Yellow

Method

Golterman, H., L. Clymo & M. A. M.

Ohnstad, 1978.)



References

Chaney, A. L. and Morbach, E. P., 1982. Modified reagents for the determination of urea and ammonia. Clin. Chem. 8, 130-132.

Golterman, H., L. Clymo & M. A. M. Ohnstad, 1978. Methods for chemical and physical analyses of freshwaters. 2nd edition. Blackwell Scientific Publishers, Oxford.

Jespersen, A-M. & K. Christophersen, 1987. Measurements of chlorophyll a from phytoplankton using ethyl alcohol as extraction solvent. Arch.Hydrobiol. 109:

445-454.

Mackereth, F.J., H. J. Heron & J. F. Talling, 1978. Water analyses: some methods for limnologists. Freshwater Biological Assoc. Scientific Publication No: 36.

Standard Methods, 22. Edition. American Health Association, 1996.

San++ Automated Wet Chemistry Analyzer, Skalar Analytical,B.V., Breda, The Netherlands.

5.8.8 Netherlands Institute of Ecology (NIOO)

Parameter


Reference

Comments

Manual Automated

Alkalinity X Si Analytics Titroline-7000, Titrasoft software 3.1 Si Analytics,

Carbon

Shimadzu TOC-L

Shimandzu Benelux, Den Bosch, The

Netherlands

DOC/ DIC X

TOC/ TIC

Chlorophyll a X HPLC ultimate 3000

Nitrogen



NH3 X Quaatro method 541502714000

Quaatro Applications, Beun de Ronde,

Abcoude, The Netherlands

NO2 X Quaatro method 5415028714100



NO3 X Quaatro method 5415028714100



TN X Flash CN analyser Interscience, Breda, The Netherlands.

Phosphorus

TP/ TDP X Quaatro method Q-031-04 Rev. 1



Silica X Quaatro method Q-038-04 Rev 1



5.8.9 Umweltbundesamt (UBA)

Parameter


Reference

Comments

Manual Automated

Alkalinity X Acidic titration by robotic titrosampler

Endpoints: DIN EN ISO 9963-1, optional:

Gran-Plot-titration acc. Sigg & Stumm

1989

Coupled with major

anion and cation

analyses (TitrIC-System)

(incl. ion mass balance)



Carbon

DOC/ DIC

TOC/ TIC X

Combustion and IR-detection by through-flow or

automatic sampler DIN EN 1484

Chlorophyll,

total (chl-a +

pheophytin) X

Photometric detection after cell disruption and

subsequent hot ethanol extraction

Endpoints: Total chlorophyll acc. Parsons

& Strickland 1963, chl a + pheophytin acc.

DIN 38412-16

Ultrasonic cell

disruption

Filterable dry

matter X Gravimetric DIN 38409-2

Major

anionic

components:

F, Cl, Br, SO4,

PO4, NO2,

NO3 X

IC-automated detection by electric conductivity,

incl. inline dialysis, chemical and CO2-surpression DIN EN ISO 10304-1

Coupled with titration

and major anion



Major

cationic

components:

Li, Na, K, Mg,

Ca, NH4 X

IC-automated detection by electric conductivity

incl. inline dialysis DIN EN ISO 14911

Coupled with titration

and major anion



Nitrogen

NH3 X

CFA-based photometric detection, indophenol blue

method (Berthelot’s reagent)

Chaney & Marbach 1962, Bucur et al.

2006, DIN EN ISO 11732, Skalar Kat Nr.

155-002w/r



NO2 X

CFA-based photometric detection, diazotization of

NO2 (Gries-Ilosvay reaction) with sulphanilamide

Bendschneider & Robinson 1952, acc. to

DIN EN ISO 13395, device specific: Skalar:

Katnr. 461-031

NO3 X


nitrite by use of coppered Cd-granules and

detection as nitrite (see above)

Wood et al. 1967, Nydahl 1976

TN/TDN

X

Pressure digestion by use of persulfate oxidation to

nitrate and subsequent detection as nitrite (see

above)

Koroleff 1983b

Joint digestion of

nitrogen + phosphor

feasible

Phosphorus

SRP X


molybdenum blue complex by use of ascorbic acid

and antimonyl tartrate

Murphy & Riley 1962, Walinga et al. 1995,

acc. to DIN EN ISO 15681-2: Skalar: Kat Nr.

503-010w/r, a + b

Optional: Low-level-

phosphate module used

below 2 µg/L PO4-P

TP/ TDP X

Pressure digestion by use of persulfate oxidation

and subsequent photometric detection as

phosphate (SRP) (see above)

Koroleff 1983a

Joint digestion of

nitrogen + phosphor

feasible

Silica X


molybdenum blue complex by use of ascorbic acid

Mullin & Riley 1955, DIN EN ISO 16264,

Skalar: Katnr. 563-052

CFA-based photometric

detection, reduction to

molybdenum blue

complex by use of

ascorbic acid



References

Bendschneider, K., Robinson, R.J. (1952): A new spectrophotometric method for the determination of nitrite in sea water. J. Mar. Res. 11, 97-96.

Bucur, B., Catala Icardo, M., Martinez Calatyud, J. (2006): Spectrometric determination of ammonia by an rFIA assembly. Revue Roumaine de Chimie 51, 101-108.

Chaney, A.L., Marbach, E.P. (1962): Modified reagents for determination of urea and ammonia. Clin. Chem. 8, 130-132.

DIN 38409-2: Summarische Wirkungs- und Stoffkenngrößen (Gruppe H), Bestimmung der abfiltrierbaren Stoffe und des Glührückstandes (H 2).

German standard methods for the examination of water, waste water and sludge; parameters characterizing effects and substances (group H); determination of

filterable matter and the residue on ignition (H 2).

DIN 38412-16: Testverfahren mit Wasserorganismen (Gruppe L), Bestimmung des Chlorophyll-a- Gehaltes von Oberflächenwasser (L16)

German standard methods for the examination of water, waste water and sludge; test methods using water organisms (group L); determination of chlorophyll a

in surface water (L 16)

DIN EN 1484: Wasseranalytik - Anleitungen zur Bestimmung des gesamten organischen Kohlenstoffs (TOC) und des gelösten organischen Kohlenstoffs (DOC);

Deutsche Fassung EN 1484-1997

Water analysis - Guidelines for the determination of total organic carbon (TOC) and dissolved organic carbon (DOC)

DIN EN ISO 11732: Wasserbeschaffenheit - Bestimmung von Ammoniumstickstoff - Verfahren mittels Fließanalytik (CFA und FIA) und spektrometrischer Detektion

(ISO 11732:2005); Deutsche Fassung EN ISO 11732: 2005.

Water quality - Determination of ammonium nitrogen - Method by flow analysis (CFA and FIA) and spectrometric detection (ISO 11732:2005)

DIN EN ISO 13395: Wasserbeschaffenheit. Bestimmung von Nitritstickstoff, Nitratstickstoff und der Summe von beiden mit der Fließanalytik (CFA und FIA) und

spektrometrischer Detektion (ISO 13395: 1996). Deutsche Fassung EN ISO 13395: 1996.

Water quality - Determination of nitrite nitrogen and nitrate nitrogen and the sum of both by flow analysis (CFA and FIA) and spectrometric detection (ISO 13395:

1996)



DIN EN ISO 15681-2: Wasserbeschaffenheit. Bestimmung von Orthophosphat und Gesamtphosphor mittels Fließanalytik (FIA und CFA). Teil 2: Verfahren mittels

kontinuierlicher Durchflussanalyse (CFA) (ISO 15681-2: 2003). Deutsche Fassung EN ISO 15681-2: 2004.

Water quality - Determination of orthophosphate and total phosphorus contents by flow analysis (FIA and CFA) - Part 2: Method by continuous flow analysis

(CFA) (ISO 15681-2: 2003)

DIN EN ISO 16264: Wasserbeschaffenheit. Bestimmung löslicher Silicate mittels Fließanalytik (FIA und CFA) und photometrischer Detektion (ISO 16264:2002).

Deutsche Fassung EN ISO 16264: 2004.

Water quality - Determination of soluble silicates by flow analysis (FIA and CFA) and photometric detection (ISO 16264: 2002)

DIN EN ISO 9963-1: Wasserbeschaffenheit-Bestimmung der Alkalinität, Teil 1: Bestimmung der gesamten und der zusammengesetzten Alkalinität (ISO 9963-1):

1994, Deutsche Fassung EN ISO 9963-1: 1995

Water quality - Determination of alkalinity - Part 1: Determination of total and composite alkalinity (ISO 9963-1: 1994)

DIN EN ISO 10304-1: Wasserbeschaffenheit- Bestimmung von gelösten Anionen mittels Flüssigkeits- Ionenchromatographie, Teil 1: Bestimmung von Bromid,

Chlorid, Fluorid, Nitrat, Nitrit, Phosphat und Sulfat (ISO 10304-1:2007), Deutsche Fassung EN ISO 10304-1:2009

Water quality - Determination of dissolved anions by liquid chromatography of ions - Part 1: Determination of bromide, chloride, fluoride, nitrate, nitrite,

phosphate and sulfate (ISO 10304-1:2007)

DIN EN ISO 14911: Wasserbeschaffenheit- Bestimmung der gelösten Kationen Li+, Na+, NH4+ , K+, Mn2+, Ca2+, Mg2+, Sr2+ und Ba2+ mittels

Ionenchromatographie, Verfahren für Wasser und Abwasser (ISO 14911:1998), Deutsche Fassung EN ISO 14911: 1999

Water quality - Determination of dissolved Li⁺, Na⁺, NH₄⁺, K⁺, Mn²⁺, Ca²⁺, Mg²⁺, Sr²⁺ and Ba²⁺ using ion chromatography - Method for water and waste water (ISO

14911: 1998)

Koroleff, F. (1983a): Determination of phosphorus by acid persulphate oxidation. - In Grasshoff, K. (ed.): Methods of sea water analysis (2nd ed.), p. 134-136.

Weinheim: Verlag Chemie.

Koroleff, F. (1983b): Determination of total and organic nitrogen after persulphate oxidation. - In Grasshoff, K. (ed.): Methods of sea water analysis (2nd ed.), p.

164-168. Weinheim: Verlag Chemie.



Mullin, J.B., Riley J.P. (1955): The colorimetric determination of silicate with special reference to sea and natural waters. - Anal. Chim. Acta 12, 162-176.

Murphy, J., Riley J.P. (1962): A modified method for the determination of phosphate in natural waters. - Anal. Chim. Acta 27, 31-36.

Nydahl, F. (1976): On the optimum conditions for the reduction of nitrate to nitrite by cadmium. Talanta 23: 349-357.

Parsons T.R. & Strickland J.D.H. (1963): Discussion of spectrophotometric determination of marine-plant pigments, with revised equations for ascertaining

chlorophylls and carotenoids. J. Mar. Res. 21: 155-63.

Skalar Kat Nr. 155-002w/r, Analyse: Ammonium, Bereich: 2 - 100 ppb P, Matrix: Abwasser. - Issue 102197/MH/97202624 97-0411.

Skalar Kat Nr. 461-031, Analyse: Nitrat + Nitrit, Meßbereich: 2 - 100 ppb P, Matrix: Seewasser. - Issue 0690899/MH/99207147. 990728 Institut

für Meereskunde

Skalar Kat Nr. 503-010w/r, a: Analyse: Phosphat, Meßbereich: 2 - 100 ppb P, Matrix: Seewasser. - Issue 060899/MH/99207147.

Skalar Kat Nr. 503-010w/r, b: Analysis: Phosphate, Range: 5 - 100 ppb P, Matrix: Surface- & drinking water. - Issue 0690803/MH/99226559.

Skalar Kat Nr. 563-052: Analyse: Silikat, Meßbereich: 0.02 - 1 ppm Si, Matrix: Wasser. - Issue 111397/MH/97202944 97-0541.

Sigg, L. & Stumm, W. (1989): Aquatische Chemie. - 396 S., 128 Abb., 37 Tab. Zürich: Verlag der Fachvereine 1989.

Walinga, I., Van Der Lee, J.J., Houba, V.J.G., Van Vark. W & Novozamsky, I. (1995): 1.7.2 Determination of phosphorus by colorimetry (automated, by flow

analyzer). - In: Walinga, I., Van Der Lee, J.J., Houba, V.J.G., Van Vark. W & Novozamsky, I. (eds.) (1995): Plant analysis manual: PANA-A1/34 - 39. Dordrecht:

Kluwer.

Wood, E.D., Armstrong, F.A.J. & Richards, F.A. (1967): Determination of nitrate in sea water by cadmium-copper reduction to nitrite. J. mar. biol. Ass. U.K. 47, 23-

31.



5.8.10 University of Helsinki, Tvärminne Zoological Station (UH)

Parameter


Reference

Comments

Manual Automated

Carbon

DOC X

High-temperature catalytic oxidation and

infrared detection

Grasshoff et al. (1999) (chapter

15)

POC X


mass spectrometric detection


17)

Chlorophyll a X

Fluorometric detection after ethanol

extraction

Baltic marine environment

protection commission (1988)

Nitrogen

NH3 X Indophenol Blue Method

Modification of Grasshoff et al.

(1999) (chapter 10.2.10);

Grasshoff (1976) (chapter 9.2)

Dichloroisocyanuric acid

as hypochlorite donor

NO2 X Automated Colorimetric Method


(1999) (chapter 10.2.8);


NO3 X

Automated Colorimetric Method, Vanadine

Chloride Reduction Method


(1999) (chapter 10.2.9);




PON X


chemiluminescence detection


17)

TDN

X


chemiluminescence detection


15)

TN X

Persulfate Digestion, Vanadine Chloride

Reduction Method


(1999) (chapter 10.2.16);

Grasshoff (1976) (chapter 9.8.3)

Phosphorus

SRP X Automated Ascorbic Acid Method


(1999) (chapter 10.2.5);


PP

X

Dry Ashing, Ascorbic Acid Method

Solorzano 1980a; Modification

of Grasshoff et al. (1999)

(chapter 10.2.12); Grasshoff

(1976) (chapter 9.1.2)

TP X

Persulfate Digestion, Automated Ascorbic

Acid Method


(1999) (chapter 10.2.13);


Silica X Automated Molybdosilicate Method


(1999) (chapter 10.2.11);




References

Guidelines for the Baltic monitoring programme for the third stage (1988). Part D. Biological determinands. Baltic Sea Environment Proceedings No. 27 D. Baltic

Marine Environment Protection Commission – Helsinki Commission.

Grasshoff, K. (Ed.) (1976). Methods of seawater analysis. Verlag Chemie.

Grasshoff, K., Kremling, K., & Ehrhardt, M. (Eds.). (1999). Methods of seawater analysis. John Wiley & Sons.

SOLORZANO, L., SHARP, J. H. (1980a). Determination of total dissolved phosphorus and particulate phosphorus in natural waters. Limnol. Oceanogr., 25(4), 754-

758.

5.8.11 University of Bergen (UIB)

Parameter


Reference

Comments

Manual Automated

Carbon

POC X Flash elemental analyses Pella & Colombo 1973

TOC1 X

High temperature catalytic

oxidation Børsheim 2000

Chlorophyll a X

Fluorometric Detection after

Acetone Extraction Parsons et al. (1984)

X

Fluorometric Detection after

Methanol Extraction Holm-Hansen and Riemann (1978)



Nitrogen

NH4+ X

Ortho-phthaldialdehyde

fluorometric Method (OPA)

Holmes et al. 1999

Adapted for microwell plate according

to Poulin & Pelletier 2007

NO3 X2

Cadmium reducing column

(nitrate to nitrite)

Parsons et al. (1992) adapted to an

autoanalyzer (San1 Segmented Flow

Analyser, Skalar Analytical B.V., The

Netherlands) as described in Rey et al.

(2000).

PON X Flash elemental analyses Pella & Colombo 1973

Phosphorus

SRP X

Ascorbic Acid - molybdenum

- tartrate blue complex Koroleff 1983

Salinity X

Direct measurement with

SAIV SD204 CTD -

Silicate

Ascorbic Acid - Molybdenum

– Oxalic - Blue Complex Valderrama 1995

1 No routine analysis

2 Performed by Institute of Marine Research (IMR) in Bergen



References

Børsheim, K. Y. 2000. Bacterial production rates and concentrations of organic carbon at the end of the growing season in the Greenland Sea. Aquat. Microb.

Ecol. 21: 115– 123. doi:10.3354/ame021115

Holm-Hansen O, Riemann B (1978). Chlorophyll a determination: improvements in methodology. Oikos 30: 438-447.

Holmes RM, Aminot A, Keroul R, Hooker AH, Peterson BJ (1999). A simple and precise method for measuring ammonium in marine and freshwater ecosystems.

Aquat Sci 56: 1801-1808.

Koroleff F (1983). Determination of nutrients. In: Grasshoff K, Ehrhardt M, Kremling K (eds). Methods in seawater analyses. Verlag Chemie: Weinheim/Deerfield

Beach, Florida. pp 125-131.

Parsons, T. R., Y. Maita, and C. M. Lalli. 1984. A manual of chemical and biological methods for seawater analysis, p.

Pergamon Press.

Pella E, Colombo B (1973). Study of carbon, hydrogen and nitrogen determination by combustion-gas chromatography. Microchimica Acta 61: 697-719.

Rey, F., T. T. Noji, and L. A. Miller. 2000. Seasonal phytoplankton development and new production in the central Greenland Sea. Sarsia 85: 329–344. doi:10.1080/

00364827.2000.10414584

Valderrama JC (1995). Methods of nutrient analysis. In: Hallograeff GM, Anderson DM, Cembella AD (eds). Manual of harmful marine microalgae. IOC manuals

and guides. UNESCO: Paris. pp 262-265.



5.8.12 Umea Marine Science Center (UMF)

Parameter


Reference

Comments

Manual Automated

Alkalinity X Potentiometric Titration

SS-EN ISO 9963-

1:1994 modified

HC-B-B151

Carbon

DOC X

High temperature combustion with NDIR

detection

HC-C-C21 / SS-EN

1484 ed. 1

modified

Chlorophyll a X Spectrofluorometry, ex 433nm/em 673nm ICES / HC-C-C21 Ethanol extraction

C and N X

Elemental analysis: High temperature

combustion with IR-detection

LECO

Corporation5

Nitrogen

NH4 X CFA (QuAAtro, Autoanalyzer ) ”Grasshoff”2 Photometric Phenol method



NO2 X CFA (QuAAtro, Autoanalyzer ) ”Grasshoff”2

Photometric

sulfanilamide/Ethylenediamine

NO3 X CFA (QuAAtro, Autoanalyzer ) ”Grasshoff”2

Photometric CD-reductor,


TN X

Simultaneous N and P Oxidative digestion

with peroxodisulfate using the borate buffer

system followed by CFA (QuAAtro,

Autoanalyzer ). ”Grasshoff”2



TDN X







Oxygen X Winkler Titration SS-EN 25813:1992

pH

HC-B-B141 / SS-EN

ISO 10523:2012 pH 7 – 10

Phosphorus

PP X Photometric Dry combustion

Solarzano3 S-EN

ISO 6878:20054

Photometric Molybdate-

ascorbic acid



TP X






ascorbic acid

TDP X






ascorbic acid

Silica X CFA (QuAAtro, Autoanalyzer) ”Grasshoff”2

Photometric Molybdate-Oxalic

acid

References

1HELCOM Combined Manual for Marine Monitoring (2015) Letters B or C refers to actual part and annex

2K. Grasshoff et al, Methods of Seawater Analysis, 2nd edition, Verlag Chemie, 1983, page 125-187; 347-376

3L.Solarzano, J. H. Sharp, Limnol. Oceanogr.., 25(4) 1980 754-758,

4SS-EN ISO 6878:2005, Water quality—Determination of phosphorus - Ammonium molybdate spectrometric method

5LECO Corporation, Thru Spec CHN/CHNS Micro Carbon/Hydrogen/Nitrogen/Sulfur Determinators. Instruction Manual Version 2.7X. Part Number 200-716, July

2015



5.8.13 WasserCluster Lunz (WCL)

Parameter


Reference

Comments

Manual Automated

Alkalinity X Titration Schwoerbel (chapter 1.2.5)

Carbon

DOC/ DIC X Oxidation and Conductometric Detection U.S. Patent No. 5,132,094

TOC/ TIC X Oxidation and Conductometric Detection U.S. Patent No. 5,132,094

Chlorophyll a X Fluorometric Detection after Acetone Extraction Arar 1997 (EPA Method 445.0)

No cell disruption and

acidification is applied

Nitrogen

NH3 X Indophenol Blue Method Ivancic & Deggobis 1984

X Indophenol Blue Method ISO 7150


X Colorimetric Method ISO 13395:1996


X Sodiumsalicylate Method Schwoerbel (chapter 1.2.14)

TN

X Persulfate Digestion, Hydrazine Reduction Method

Clesceri 1999 (4500-P, chapter J);

ISO 13395:1996

Phosphorus

SRP X Ascorbic Acid Method Grasshoff 1999 (chapter 10.2.5)

X Automated Ascorbic Acid Method ISO 15681-2



PP

X

Dry Ashing and Ascorbic Acid Method

Solorzano 1980a; Grasshoff 1999 (chapter

10.2.5)

Digestion modified from

Solorzano

TP/ TDP X Persulfate Digestion and Ascorbic Acid Method Grasshoff 1999 (chapter 10.2.13)

Silica X Molybdosilicate Method or Heteropoly Blue Method Clesceri 1999 (4500-SiO2, chapter C or D)

References

Arar, E. J., & Collins, G. B. (1997). Method 445.0: In vitro determination of chlorophyll a and pheophytin a in marine and freshwater algae by fluorescence.

Cincinnati: United States Environmental Protection Agency, Office of Research and Development, National Exposure Research Laboratory.

Clesceri, L. S., Greenberg, A. E., & Eaton, A.D. (1996). Standard methods for the examination of water and wastewater. APHA, AWWA and WPCF, Washington DC.

DIN EN ISO 15681-2, Water quality - Determination of orthophosphate and total phosphorus contents by flow analysis (FIA and CFA) - Part 2: Method by

continuous flow analysis (CFA), 06-2001

Grasshoff, K., Kremling, K., & Ehrhardt, M. (Eds.). (1999). Methods of seawater analysis. John Wiley & Sons

ISO 7150-1: 1984, Water quality—Determination of ammonium, manual spectrometric method

ISO 13395:1996, Water quality—Determination of nitrite nitrogen and nitrate nitrogen and the sum of both by flow analysis (CFA and FIA) and spectrometric

detection.

Ivančič, I., & Degobbis, D. (1984). An optimal manual procedure for ammonia analysis in natural waters by the indophenol blue method. Water Research, 18(9),

1143-1147.

R. Godec, et. al., “Method and apparatus for the determination of dissolved carbon in water” U.S. Patent No. 5,132,094)

Solorzano, L., & Sharp, J. H. (1980). Determination of total dissolved phosphorus and particulate phosphorus in natural waters. Limnology and Oceanography,

25(4), 754-7



5.9 References 5 – Water Chemistry

[1] G. WOLFRAM, K. DONABAUM, and M. T. DOKULIL,

“Guidance on the Monitoring of the Biological Quality

Elements Part B2 - Phytoplankton,” Vienna, 2015.

[2] WURL, Oliver (Hg.). Practical guidelines for the analysis of

seawater. CRC press, 2009.

[3] NOLLET, Leo ML; DE GELDER, Leen SP (Hg.). Handbook of

water analysis. CRC press, 2000.

[4] WHO, Handbook: Good Laboratory Practice (GLP): Quality

practices for regulated non-clinical research and

development, 2nd ed. Switzerland: TDR/WHO, 2009.



[6] Council Directive, “89/391/EEC of 12 June 1989 on the

introduction of measures to encourage improvements in the

safety and health of workers at work,” Off. J. Eur.

Communities, vol. 183, no. 29, p. 8, 1989.

[7] OSHA, “European directives on safety and health at work

- Safety and health at work.” [Online]. Available:

https://osha.europa.eu/en/safety-and-health-

legislation/european-directives. [Accessed: 16-Jun-2017].

[8] European Commission, “EU Waste Legislation.” [Online].

Available:

http://ec.europa.eu/environment/waste/legislation/a.htm.

[Accessed: 16-Jun-2017].

[9] European Commission, “COMMISSION DECISION of 3 May

2000 replacing Decision 94/3/EC establishing a list of wastes

pursuant to Article 1(a) of Council Directive 75/442/EEC on

waste and Council Decision 94/904/EC establishing a list of

hazardous waste pursuant to Article 1(4) of Council Directive

91/689/EEC on hazardous waste (notified under document

number C(2000) 1147) (Text with EEA relevance)

(2000/532/EC),” Off. J. Eur. Communities, vol. 69, 2000.


D3.1-4: guidance document on sampling, analysis and

counting standards for phytoplankton in lakes,” pp. 1–51,

2012.

[11] UNESCO/IOC, MICROSCOPIC AND MOLECULAR

METHODS FOR QUANTITATIVE PHYTOPLANKTON ANALYSIS.

Paris: O. (IOC Manuals and Guides, no. 55.)

(IOC/2010/MG/55), 2010.

[12] Aarhus Universitet – National Environmental Research

Institute (AU) and Middle East Technical University (METU),

“EU FP7 244121, REFRESH Deliverable 3.4: Protocols for field

experiments and monitoring along a climate gradient in task

2,” 2014.

[13] GRASSHOFF, Klaus; KREMLING, Klaus; EHRHARDT,

Manfred (Hg.). Methods of seawater analysis. John Wiley &

Sons, 2009.

Co-funded by the European Union D4.1 Standard Operating Procedures| 6 High Frequency Monitoring


6. High Frequency Monitoring


Calibration To standardize or correct sensors after determining, by measurement or comparison

with a standard, the correct value [1].

Drift The lack of repeatability caused by fouling of the sensor, shifts in the calibration of the

system, or slowly failing sensors [2].

Deployment The way that the sensor comes into contact with the ambient water [3].

Multiprobe The combination of several sensors, electrodes, or probe assemblies into a complete,

stand-alone piece of equipment which simultaneously measures several parameters for

profiling, spot-checking, or logging readings and data.

Long-term drift The slope of the regression line derived from a series of differences between reference

and measurement values obtained during field testing expressed as a percentage of the

working range over a 24 h period [4].

Probe A small tube containing the sensing elements of electronic equipment. It is an essential

part of the water quality monitoring system since it obtains measurements and data

which can be stored, analyzed, and eventually transferred to a computer.

Profiling Lowering a probe through a water column to characterize the vertical distribution of

parameters.

Sensor The fixed or detachable part of the instrument that measures a particular field

parameter [5].

Short-term drift The slope of the regression line derived from a series of measurements carried out on

the same calibration solution during laboratory testing and expressed as a percentage

of the measurement range over a 24 h period [4].



6.2 Equipment and supplies

Equipment used in automated sampling are (Adopted from (Canadian Council of Ministers of the

Environment, 2011)):

1. the sensors (electrical, electrochemical, or optical) used to collect the data, which respond to

changing water conditions with an output signal that is processed, displayed and recorded. The

choice of the sensor depends on the parameters, the required specifications, the operating

conditions, and required lifespan.

2. the accessory equipment:

✓ a data logger which may be contained within a (multi-) probe or connected externally. Data

filtering and processing are completed within the data loggers. The time interval of the

recorded samples is determined by the user. The duration of individual samples is a function

of the sensors.

✓ power supply can be internal batteries (which are contained within the sensor), external

batteries, which should be a good quality gel-cell type, or a deep discharge sealed lead-acid

style; and solar panels (used for satellite transmission).

✓ Best Practice Advice: Residential (220V) and solar power sources can be used as

auxiliary power to the primary battery for recharge purposes. Residential and solar

power sources should not be directly connected to an instrument, as voltage spikes

can occur and cause the entire system to fail. Use of a voltage regulator is

recommended when connecting an auxiliary power source to the primary battery.

✓ a means of retrieving the data; Communication and data retrieval can be done on-site with

a laptop or hand-held display. Data retrieval can also be achieved remotely in real-time

using phone or satellite communication.

3. the cables and adapters, (instrument and site-specific) to connect the external batteries to the

sensor or the solar panel to the external battery.

6.3 Health and safety indications

✓ Please see the Water Chemistry SOP for working and personal protection equipment suggested for

use in water sampling.

✓ Conductivity standard solutions, pH buffers, DO and pH reference solutions are nontoxic but can

irritate eyes and other sensitive areas because of their high salt content.

✓ Rhodamine dye; used as a calibrant for chlorophyll-a measurements, is also nontoxic, but stains

everything it contacts.



6.4 Field Parameters and Sensors

Sensors can be in the form of individual instruments or as a single instrument including different sensors with

many combinations of field parameters (multi probes) [5].

In this section, the information on the field parameters (included in this SOP) that can be measured with

automated sensors will be explained in detail. In addition, best practice advice will be provided for

measurements.

6.4.1 Temperature

Temperature is a critical environmental factor affecting physiological processes in organisms, the density of

water, solubility of constituents (such as oxygen in water), pH, conductivity, and the rate of chemical

reactions in the water [6], [7]. Monitoring of temperature thus is mandatory in any outdoor experiment.

Depending on the fluctuations, and whether temperature itself is a treatment factor, the frequency of

monitoring must match the experimental design. The monitoring of temperature along a vertical profile

within a mesocosm provides information on the degree of stratification or homogenization of the water

column [7].

Among many temperature sensors, thermistors are well adapted to measure the natural water temperature

in mesocosms [7]. Thermistors that are made of a solid “semiconductor having resistance that changes with

temperature” are incorporated into digital thermometers, other parameter instruments such as conductivity

and pH meters and/or multi-parameter instruments [5]. The thermistor thermometer converts changes in

resistance into temperature units. The resistance varies inversely with temperature, but, more importantly,

their baseline resistance value and coefficient of variation are generally large. In some applications, an

improvement can be reached with a software correction for minimization of linearization errors.

✓ The measurement range of temperature sensors are wide (-40 to +95°C, commonly). The

measurement range of a thermistor used in a high-frequency monitoring station in a recent

mesocosm study was from 0 to 40 °C [7].

✓ Thermistors are one of the most accurate types of temperature sensors. The accuracy of thermistors

is from 0.01 to 0.02°C in a recent mesocosm study [7].

✓ Thermistors require little maintenance and are relatively inexpensive.

✓ Diel fluctuations in water temperature tend to follow the fluctuations of atmospheric temperature

[7].

✓ Many other parameters measured by automated probes (conductivity, DO, pH) are affected by

temperature. In a multiprobe, wrong calibration of the temperature probe affects multiple

parameters.



6.4.2 pH

pH is the mathematical notation defined as “the negative base-ten logarithm of the hydrogen-ion activity,

measured in moles per liter of a solution” [8, p. 3]. It is a primary factor governing the chemistry of natural

water systems that affect physiological functions of plants and animals.

✓ The amount of dissolved gases, such as carbon dioxide, hydrogen sulfide, and ammonia affect pH

level of water.

1) The instrument system that is commonly used to measure pH consists of a pH electrode; gel-filled or

liquid-filled, which is a special type of ion-selective electrode (ISE), designed specifically for the

measurement of the hydrogen ion (summarized from [8]). The pH electrodes have “a glass

membrane, a reference and a measurement electrode, ionic (filling) solution, and a reference

junction” [8, p. 6]. Sensors used in submersible monitors typically are combination electrodes in

which a proton (H+

)-selective glass-bulb reservoir is filled with an approximate pH-7 buffer.

✓ A clean, undamaged glass membrane is necessary for performing an accurate measurement

of pH.

✓ Remember to check the concentration of the filling solution as any change in concentration

level will result in sensitivity loss.

✓ Remember to check that the junction on the pH electrode is not clogged; a clogged electrode

may not function properly.

6.4.3 Dissolved Oxygen (concentration and saturation)

Sensors of dissolved oxygen (DO) do not measure oxygen in milligrams per liter or parts per million but, the

actual sensor measurement is proportional to the ambient partial pressure of oxygen; which can be displayed

either as percent saturation or in milligrams per liter, depending on user input. The concentration of

dissolved oxygen is an output of the calculations based on temperature and salinity of water [9]. The

atmospheric air pressure is another factor in changing the concentration of DO of the water.

✓ Since salinity and water temperature has an impact on the calculation of the dissolved O2

concentration, DO sensors should be associated with a temperature and a conductivity or salinity

sensor via a data logger for necessary correction in real time [7].

✓ In surface waters, DO concentrations typically range from 2 to 10 milligrams per liter (mg/L).

Luminescence-based optical sensors, which are based on dynamic fluorescence quenching, have a light-

emitting diode (LED) to “illuminate a specially designed oxygen-sensitive substrate that, when excited, emits

a luminescent light witha lifetime that is directly proportional to the ambient oxygen concentration” [5].

Most of the oxygen sensors used in marine mesocosm experiments are “based on oxygen dynamic



luminescence quenching of a platinum porphyries complex”, or in short, luminescence-based optical sensors

which can be used for long-term recording of dissolved O2 due to their strength, their immunity to biofouling

and their easiness of cleaning [7].

✓ Remember that contact with organic solvents can compromise sensor integrity or performance.

● The maintenance routine and schedule for optical sensors is infrequent.

✓ Optical-sensor maintenance is recommended by manufacturer guidelines that are specific to the

type of sensor in use and the conditions to which the sensor has been subjected [9].

✓ In a mesocosm experiment, calibration is recommended at the start of the experiment (plus bi-

monthly calibration in long-term experiments). In addition, sensors should be checked for

potential drift at the end of the mesocosm experiments.

✓ The manufacturers generally recommend annual to biannual replacement of the luminophore-

containing module as well as calibration check by them.

6.4.4 Conductivity

Conductivity is “a measure of the capacity of water to conduct an electrical current” and it is also a function

of the types and quantities of dissolved substances in water [10, p. 3]. Specific conductance is the

conductivity expressed in units of micro Siemens per centimeter at 25°C (μS/cm at 25°C).

✓ As concentrations of dissolved ions increase, the conductivity of the water increases.

The water conductivity in mesocosms can be measured by two techniques: electrode cell method and

electromagnetic induction method. The first method is commonly used for measuring low conductivities,

especially in pristine environments. Use of the second method can be preferred for high conductivity

measurements, especially in marine mesocosms [7].

Based on the methods, conductivity sensors generally are of two types, contact sensors with electrodes and

sensors without electrodes. Specific conductance sensors with electrodes require the user to choose a cell

constant (the distance between electrodes (in centimeters) divided by the effective cross-sectional area of

the conducting path (in square centimeters) for the expected range of specific conductance. One shall

choose a cell constant on the basis of expected conductivity based on Table 6-1.

✓ The greater the cell constant, the greater the conductivity that can be measured.

✓ Conductivity is temperature dependent, hence must be monitored in combination with temperature.

Wrong calibration of the temperature probe gives biased conductivity.



Table 6-1: The cell constants for contacting-type sensors with electrodes and corresponding conductivity ranges (Adopted from [10])

Conductivity range, in microSiemens per

centimeter

Cell constant, in 1/centimeter

0.005 – 20 0.01

1 – 200 0.1

10 – 2000 1.0

100 – 20000 10.0

1000 – 200000 50.0

Electrodeless-type sensors operate by inducing an alternating current in a closed loop of the solution, and

they measure the magnitude of the current.

✓ Electrodeless sensors avoid errors caused by electrode polarization or electrode fouling as they are

immune to biofouling [7].

6.4.4.1 Salinity

Salinity is the total quantity of dissolved salts in water. It can be calculated based on conductivity

measurements, as conductivity is a tool to estimate the amount of chloride in water [5]. Salinity is most

commonly reported using the Practical Salinity Unit (PSU), described by Lewis (1980) [11] which is, a scale

developed relative to a standard potassium-chloride solution and based on conductivity, temperature, and

barometric pressure measurements [5]. Currently the salinity expressed dimensionless.

Salinity may not be directly measured but is derived from conductivity – temperature – depth/pressure

measurements. For conversion among related parameters (conductivity, salinity, temperature, and pressure)

several packages are provided within the R statistical language (e.g., marelac; seacarb).

Recently, “a compact microstrip feed inset patch sensor” has been developed for measuring the salinity in

seawater, which was claimed to bring better sensitivity to salinity changes (than commonly used sensors

using conductivity change to measure the change in salinity) [12].



6.4.5 Underwater Light (PAR)

Photosynthetically Active Radiation (PAR) is defined as radiation in the range from 400 to 700 nm,

corresponding to the spectral part of daylight used by phototrophic organisms for photosynthesis. Measuring

PAR can provide “an excellent gauge of how much light is available to photosynthetic organisms.

Upwelling and downwelling radiation are two aspects of underwater PAR. Upwelling radiation is radiation

received from below the sensor due to reflectance off a lower surface of some type, while downwelling

radiation is a measure of radiation from above the sensor, usually due to sunlight or other external light

sources.

There are mainly two types of sensors used commonly to measure underwater PAR: planar and scalar

sensors. Planar sensors have a flat light collecting surface that responds to light that impinges on their

surface from downward directions. Planar sensors tend to underestimate PAR because the collecting surface

does not absorb upwelling radiation or light that reflects off particles in the water and the sediment surface

[13]. On the other hand, scalar PAR sensors have a hemispherical or spherical collecting surface that

functions to absorb light from 2p to 4p steradians, (sr, i.e., Standard International (SI) unit of solid angular

measure) respectively. They record more accurate measurements of total underwater PAR [13] as they

absorb diffuse radiation from most directions [14].

✓ According to Long and colleagues (2012), “cosine-corrected planar sensor”, which is another sensor

for PAR measurements; will produce more accurate measurements of PAR than a planar sensor

without cosine correction especially under light conditions which are not ideal for accurate results

such as during sunrise and sunset [14, p. 417].

✓ The planar sensors are insufficient for studies, for instance, involving phytoplankton residing within

the water column where diffuse radiation may be a significant form of available light [14]. Instead,

spherical sensors should be used for quantification of light in the water column.

✓ Advantages and disadvantages of all three types of sensors (planar, cosine-corrected planar and

scalar sensors) are summarized in Table 6-2 below (Adopted from [14]).



Table 6-2: The advantages and disadvantages of the three different instruments that can be used in PAR measurements (Adopted from [14]). Note for quantification of light in the water column, ideally spherical sensors are used (planar sensors will only capture e.g. the incoming light, depending on their exposition in the water column).

Sensor type Advantages Disadvantages

Planar Sensor (ex.

HOBO pendant

logger)

✓ Inexpensive

✓ Simple field deployment

✓ Temperature sensor

✓ Use of multiple data loggers,

reduction in data loss

✓ Small, easy to handle and mount

✓ Can be used for microscale

measurements

✓ Average out variations with

multiple loggers

✗ Data requires heavy post-processing

✗ Limited data logging period

✗ Light intensity sensor, measuring in

the unit LUX, rather than PAR

✗ Records at user-specified intervals

(no integration)

✗ No stability or accuracy reported by

the manufacturer

✗ Housing scratches and degrades

easily, shading sensor

Cosine-corrected

planar sensor (ex.

Odyssey

Integrating PAR

sensor

✓ Fairly inexpensive

✓ Simple field deployment

✓ User-specified integration periods

✓ Small, easy to handle and mount

✗ Difficult to download/open data if

using often

✗ Needs to be calibrated

✗ Difficult battery replacement

✗ No stability/accuracy reported by

the manufacturer

Spherical Sensor

(ex. LICOR LI-

193SA)

✓ Guaranteed factory calibration (+-

5%)

✓ User-specified integration periods

✓ Excellent angular response,

stability, sensitivity

✗ Expensive

6.4.6 Turbidity

Turbidity is “an expression of the optical properties of a liquid that causes light rays to be scattered and

absorbed rather than transmitted in straight lines through a sample” [15]. Turbidity is caused by the presence

of suspended and dissolved matter, such as clay, silt, finely divided organic matter, plankton, and other

microscopic organisms, organic acids, and dyes [15]. Color, either of dissolved materials or of particles

suspended in the water can also affect turbidity [5]. The most common unity of turbidity is Nephelometric

Turbidity Units (NTU).

Turbidity indicates the PAR availability in the water column of the mesocosm. Turbidity is typically measured

at a wavelength near 850-880 nm (documentation turner sensors).



✓ Note that some combined chl-a & turbidity sensors measure turbidity at 700 nm for correcting chl-a

for background turbidity (Seabird documentation).

Numerous methods and instruments can be used to measure turbidity. Because different measurement

technologies result in different sensor responses, the available turbidimeters are categorized according to

the instrument design as follows:

1. Type of incident light source (incandescent, LED, laser)

2. The detection angle (90°, 180° (attenuated angle), 0 - 45° (backscatter angle))

3. Number of the scattered/ attenuated light detectors used (multiple detection angles, dual light

source-detector)

The nature of particles could affect turbidity readings with 0.5-1.0 NTU. In long-term studies, this can be up

to 10 NTU [16]. In addition, Ruzycki and colleagues (2014) reminded that the readings could be affected due

to probable discharge, color, particle size, sediment concentrations, mineral composition and organic matter

[17].

Measurement and documentation for submersible turbidity sensors discussed thoroughly in Anderson (2005)

[15].

It is important to note that, for a valid comparison of turbidity data over time, between sites, and among

projects, it is recommended to use with identical optical and data-processing configurations. Because of the

potential to generate data with a high degree of variability when different technologies are used, use of units

in reporting is recommended to express the measure of turbidity accordingly. In other words, units indicate

the type of technology used in measuring turbidity (See Table 6-3).

✓ Nephelometric (90°) near-infrared wavelength technology is used commonly, which report data in

NTU.

Table 6-3: The reporting units employed when using given instrument design (Adopted from [18]).

Instrument Design Reporting

Unit

Nephelometric non-ratio turbidimeters (NTU)

Ratio white light turbidimeters (NTRU)

Nephelometric, near-IR turbidimeters, non-ratiometric (FNU)

Nephelometric, near-IR turbidimeters, ratiometric (FNRU)

Surface scatter turbidimeters (SSU)

Formazin backscatter unit (FBU)



Backscatter unit (BU)

Formazin attenuation unit (FAU)

Light attenuation unit (AU)

Nephelometric turbidity multibeam unit (NTMU)

6.4.7 Depth

Depth is routinely measured by many multi probes as to relate other parameters to a given depth. Depth is

measured with the help of a vented or non-vented pressure sensor (ex. YSI EXO1 and EXO2). With a non-

vented sensor, “a differential strain gauge transducer measures pressure with one side of the transducer

exposed to the water and the other side exposed to a vacuum” [19, p. 17]. Then, depth is calculated from

the pressure exerted by the water column minus atmospheric pressure.

✓ In non-vented sensor, depth measurement can be influenced by barometric pressure, water density,

and temperature.

On the other hand, vented level sensors use “a differential transducer with one side exposed to the water”

[19, p. 137] (YSI, 2018, p. 137). Unlike non-vented ones, the other side of the level transducer is vented to

the atmosphere. Accordingly, the transducer will measure the water pressure exerted by the water column,

with a particular sensor that is vented to the outer atmosphere as a tube that runs through the sonde and

cable. According to the YSI handbook (2018), this tube must remain open and vented to the atmosphere,

without any distraction of foreign objects, to function appropriately (YSI, 2018) [19].

6.4.8 Chl-a (including fluorescence per algal groups and total)

Chlorophyll-a absorbs light in the blue and red parts of the visible electromagnetic spectrum. Chlorophyll

fluorescence is the red light re-emitted by chlorophyll molecules when excited by a light source [20].

Chlorophyll fluorescence is a non-invasive method for analyzing photosynthetic energy conversion of higher

plants, algae, and bacteria [20].

As previously put forward by Mostajir and colleagues (2012), “the continuous measurement of chlorophyll a

and phycoerythrin concentrations by fluorescence sensors (such as YSI EXO2 sensors) is used to monitor the

presence and the temporal dynamic of the algal bloom (chlorophyll a measurement) or cyanobacteria

(phycoerythrin measurement)” in a mesocosm experiment [7]. The measurements can also be used to

determine “the diel variations of pigment concentration (particularly chlorophyll a), which is affected by

change of phytoplankton biomass and photo acclimation processes” [7, p. 313].



In situ fluorometers with an excitation (light) source, such as a Xenon lamp, laser or LED, are regarded as

highly sensitive tools for the quantification and analysis of phytoplankton, which offer continuous

measurement of chlorophyll concentrations in the field. They neither require pre-treatment nor a large

sample volume [20]. They can be used with a profiler and integrated CTD sensor to collect data at different

depths of a mesocosm [21].

✓ It should be noted that the excitation (light) source can greatly affect the quality of fluorescence

signal because of its importance as a prerequisite of light-induced fluorescence measurement.

Different light sources have their own advantages and disadvantages. For a brief comparison of the

light sources used, please read Zeng and Li (2015) [20].

✓ In addition, the type of detector, which is the receiver of the fluorescence, have an important impact

on the detection limit and measurement frequency. Selecting a high-performance detector with low

noise can improve both the detection limit and the measurement accuracy [20].

✓ In-situ measurement of chl-a by autofluorescence is dependent on the physiological state of the algal

cells. In general, dark adaptation is recommended for comparable measurements. For probes that

are permanently installed for in-situ measurements in mesocosms, dark adaptation is not an option.

Here, nighttime readings may be more comparable than readings from daytime. However, as the

most classical measurements (e.g. HPLC) realized during the day, the comparison of the results could

not be possible.

6.4.9 Nitrogen (Nitrate)

The continuous nutrient concentration monitoring in real time by in situ nutrient probes allows determining

sources, sinks, and dynamics of different nutrients in natural environments and can be adapted for the

mesocosm experiment [7].

Most in situ nutrient (nitrate, nitrite, phosphate, ammonia, etc.) analyzers automatically measure dissolved

nutrients by “using wet chemical techniques of in-flow analysis based on standard laboratory analytical

methods (spectrophotometry and fluorometry)” [7, p. 311]. These analytical probes are equipped with one

or multiple reagent-delivering modules and standards, and one or multiple electro-optical detectors,

number of which depends on the measurement of one or several nutrients.

✓ The frequency of measurement is affected by the amount of reagent and standard loaded in the

probe [7].

✓ To avoid the drifts and the degradation of optics, reagents, calibration standards, and also biofouling

in the sensor sampling line, frequent maintenance is necessary, which includes reagents and

standards change, complete clean up and in situ recalibration of the probe [7]. Please see the next

section for basic calibration recommendations.



The nutrient (ammonium, nitrate, and chloride) can also be measured by ion-selective electrodes (ISEs),

which are commonly used in in situ water quality monitoring as they can be used “directly in the medium to

be tested, have a compact size, and are inexpensive” [22].

6.4.10 Multi-probes (Sensors, profilers)

Multi-probes are the sensors that allow for the continuous monitoring of several common field parameters

altogether. These sensors are available individually or bundled together into a multi-parameter sensor with

several sensors attached to a single unit [23].

✓ Note that the accuracy of the temperature probe affects multiple other parameters in multi-probes

(e.g., Oxygen, pH, conductivity).

The types and number of sensors that can be bundled in each sensor depend on the instrument model and

manufacturer. CTD sensors (Conductivity, Temperature, and Depth) are a good example of a basic

multiparameter sensor. CTDs measure pressure (rather than depth) based on the relationship between

pressure and depth, which also involves water density and compressibility as well as the strength of the local

gravity field. The output data of CTDs can be used to calculate salinity, density, and sound velocity.

Aside from temperature, conductivity and depth, dissolved oxygen, pH, PAR and turbidity sensors are

commonly bundled together into a multiprobe in mesocosm studies. YSI EXO Series, Hydrolab DS5X and

Aanderaa Seaguard series are the commonly used multiparameter sensors throughout the Aquacosm

community. YSI EXO series (ex. EXO2) allow the user to collect data from up to six user-replaceable sensors

and an integral pressure transducer (YSI) for 17 parameters (temperature, conductivity, salinity, depth,

pressure, pH, Oxygen Reduction Potential (ORP), dissolved oxygen, Turbidity, Chlorophyll a, Blue-Green Algae

(Phycocyanin and Phycoerythrin), fDOM (CDOM), Ammonium, Nitrate, Chloride, Total Dissolved Gas (TDG)

and Total Suspended Solids (TSS)). In addition, optionally, they have a bulkhead (made from titanium) port

for a central wiper (or an additional sensor) and an auxiliary port on top of the sensor. Hydrolab Datasonde

series (ex. DS5X) allow the user to collect data for 17 different parameters (temperature, conductivity, depth,

pH, Oxygen Reduction Potential (ORP), dissolved oxygen (LDO and Clark Cell), Turbidity, Chlorophyll a, Blue-

Green Algae, Rhodamine WT, Ammonium, Nitrate, Chloride, Total Dissolved Gas (TDG) and Ambient Light

(PAR)) with up to seven sensor ports. It has a central cleaning system to minimize (bio)fouling of the sensors;

DO, pH, ion-selective electrodes, chlorophyll, blue-green algae, rhodamine, and turbidity (See Figure 6-1).

Aanderaa SeaGuard Series (ex. SeaGuard String) are commonly used in marine mesocosms. The SeaGuard

String is designed to be connected to the SeaGuard String logger; which can be connected with up to 25

sensor nodes. In addition, up to 6 sensors can be mounted onto the Top-end Plate of the multiparameter

instrument.



Figure 6-1: Hydrolab DS5X instrument design, a type of multiparameter sensor, commonly used in mesocosm studies

Some researchers prefer to use the single temperature sensors together with temperature sensor bundled

into a multiparameter (ex. Testo AG; Testo 108, Campbell Scientific; Temperature probes). Other than that,

Underwater PAR sensors (LICOR; LI-192 and 193), Chlorophyll-a sensors (Sea Bird Scientific-Wetlabs; ECO

Series) and Light spectrum sensors (Ocean Optics; USB Series Spectrometers) are commonly selected to be

single parameter probes.

6.5 Maintenance of the station and the equipment

To obtain the most accurate and most complete records possible; periodic verification of sensor calibration,

maintenance of the equipment and the monitoring station, troubleshooting of sensors and data loggers (if

applicable) should be performed regularly.

Maintenance frequency

Maintenance frequency generally is governed by the study objectives as well as the biofouling rate of the

sensors, which vary by sensor type, hydrologic and environmental conditions [5].

✓ According to Wagner and colleagues (2006), the performance of temperature and specific

conductance sensors tends to be less affected by fouling than DO, pH, and turbidity sensors [5].

For a high degree of accuracy, maintenance should be weekly or more often. In nutrient-enriched

mesocosms and moderate to high temperatures, maintenance as frequently as every third day is

recommended [5]. In addition, monitoring disruptions as a result of recording equipment malfunction,

sedimentation, electrical disruption, debris, ice, pump failure, or vandalism also may require additional site

visits.



● Satellite telemetry is recommended for sensors where lost records will critically affect research

objectives. Satellite telemetry (if applicable) can be used to verify proper equipment operation on a

daily basis and can help in recognizing and correcting problems quickly [5].

Maintenance actions (adopted from [5]):

Daily maintenance acts (for sites equipped with telemetry)

✓ Daily review of sensor function and data download

✓ Battery (or power) check

✓ Deletion of spurious data, if necessary

Maintenance acts during weekly field visits

✓ Inspection of the site for signs of physical disruption

✓ Inspection and cleaning of the sensor(s) for fouling, corrosion, or damage

✓ Inspection and cleaning of the deployment tube

✓ Battery (or power) check

✓ Time check

✓ Regular sensor inspection at the field (Calibration check, explained in details in Section 6.6.1)

✓ Calibration of the field meter(s)

✓ Downloading of data

6.5.1 Regular Sensor Inspection at the Field

Sensor inspections are required to verify that a sensor is working properly. Field trips for sensor inspection

will provide “an ending point for the interval of water-quality record” since the last visit and “a beginning

point for the next interval of water-quality record” [5].

1. Record the initial sensor readings (1) in the mesocosm: The initial sensor readings (1) of the

equipment are compared to readings from a calibrated field meter, which can provide a

reasonable comparison basis and an indication of potential electronic calibration drift and fouling

errors, ideally located at the same measuring point” in the mesocosm. The initial sensor reading

will be the last data recorded since the previous inspection visit.

✓ Readings from a calibrated field meter should not be used in computations (exceptional: the

temperature sensors), they are only used to assess “cross-section variability and

environmental changes that may occur while the monitor is being serviced”, as well as to

detect the fouling and probable calibration drift (in case the environmental conditions are

stable or slowly changing) [5].



2. Clean the sensors and recording equipment (if required)

✓ The sensor should be removed from water for servicing while the field meter remains in its

place. First, the removed sensor should be inspected for any sign of fouling; such as chemical

precipitates, stains, siltation, or biological growths. If any are observed, it should be recorded

in the field notes (see Field Form) before cleaning.

✓ For all the sensors, during the cleaning process, care should be given to ensure that the

electrical connectors are kept clean and dry.

✓ The sensors (and the recording equipment) should be cleaned according to the specifications

provided by the manufacturer.

✓ Best practice advice on cleaning for each parameter is provided in Section 8 of this SOP.

3. Record the cleaned-sensor readings (2) in the mesocosm: The cleaned sensor is returned to the

water for the cleaned-sensor readings (2). The cleaned-sensor readings, field meter readings and

reading times should be recorded in the field notes.

✓ The difference between the initial sensor reading (1) and the cleaned-sensor reading (2) is

the sensor drift caused by fouling (including chemical precipitates, stains, siltation, or

biological growth).

✓ One should note that difference might not be always representative due to

change in growth or loss of organisms in time. Those should also be

considered in calculations.

4. Do the calibration checks of sensors by using appropriate calibration standards (solutions): The

sensors should be removed from water for calibration checks when all readings are recorded.

Calibration checks of the sensor are performed in calibration standard solutions. The cleaned-

sensor readings (2a) in the calibration standard solutions are recorded in the field form.

✓ During field visits, calibration of all sensors should be checked with (two) standard solutions

that bracket the range of expected environmental conditions. A third standard solution can

be used additionally near the ambient environmental conditions, before any adjustments are

made to the monitor calibration.

✓ The difference between the cleaned-sensor readings in calibration standard solutions (2a)

and the expected reading ((2) or the calibration criteria) in these solutions is the sensor

(calibration) drift error.

✓ Best practice advice on calibration for each parameter is provided in Section 6.8 of this SOP.

5. Re-calibrate the sensors if the readings are beyond the accuracy ranges: Field re-calibration

should be performed if the cleaned-sensor readings obtained during the calibration check differ

by more than the calibration criteria (See Table 6-4).



✓ Spare monitoring sensors might be needed to replace the ones that fail calibration after re-

calibration, proper maintenance by the user and troubleshooting steps have been applied.

✓ Best practice advice on troubleshooting are provided in Section 6.6 of this SOP.

6. Record the final sensor readings (3) in the mesocosm: A set of initial readings (3) should be

taken as the start of the new record.

✓ Under rapidly changing conditions (i.e., change in the parameter that exceeds the calibration criteria

within 5 minutes) or when measurements are fluctuating, the site inspection acts should be modified

accordingly. For more details on sensor inspection at rapidly changing conditions, please refer to

(Wagner and colleagues (2006) [5].

● Validation/ cross comparison: As a best practice advice, use your maintenance visits to collect other

data to support the monitoring effort: e.g. Secchi disk and water temperature readings, water

samples for chl-a extraction, nutrients, DOC, etc. For meteorological data, make an independent

measurement of air temperature and a visual check on wind direction. While these measurements

may seem unnecessary at the time they can easily be incorporated into your maintenance visits, and

they are invaluable for confirming and strengthening patterns shown in the sensor data [24].

Table 6-4: Calibration criteria for continuous measurements (variation outside the value shown requires re-calibration)

Measurement Calibration Criteria Reference Respectively

Temperature ± 0.2 ºC

± 0.3 ºC

[5], [1]

[25]

(specific) conductance ± 5 μS/cm or ±3 % of the measured value, whichever is greater

± 10% of reading

[5]

[25] & [1]

Dissolved Oxygen ± 0.3 mg/L [5], [1] & [25]

Dissolved Oxygen (%

saturation)

± 5% saturation [1]

pH ± 0.2 pH unit

± 0.5 pH unit

± 0.3 pH unit

[5]

[25]

[1]

Turbidity ± 0.5 turbidity unit or ± 5% of the measured value, whichever is greater

± 10% of range

[5]



[25]

Underwater Light (PAR) ± 3 - 5% (for quantum sensors)

Depth

Chl-a Exceeding the accuracy of the sensor (the accuracy of the sensor is

provided in the handbook provided by the manufacturer)

± 3% for signal level equivalents of 1 ppb rhodamine WT dye or higher

using a rhodamine sensor

[25]

Light spectrum Each degree of separation from the NIST-calibrated light source

introduces some uncertainty, yielding a total estimated uncertainty of

within 10% for most Ocean Optics calibration light sources (a value that

is typical for the industry).

[26]

Nitrogen Ammonium( NH4): ±10% of reading or ±2 mg/ L-N, whichever is greater

Nitrate( NO3): ±10% of reading or ±2 mg/ L-N, whichever is greater

[19]

(±, plus or minus value shown; °C, degree Celsius; μS/cm, microsiemens per centimeter at 25 °C; %, percent; mg/L,

milligram per liter; pH unit, standard pH unit; turbidity unit is dependent on the type of meter used)

6.5.2 Best Practice Advice on Sensor-specific Field Cleaning and Calibration

6.5.2.1 General Considerations

✓ The manual of the manufacturer should be the main source of information for calibration. For regular

calibration frequency at the field site, manufacturer’s advice should be considered. Besides,

calibrating individual sensors or multi-parameter sensors at least once a month is a common good

practice advice. If recommended differently than this SOP, the manufacturer’s guidelines must be

followed for both single and multi-parameter sensors.

6.5.2.2 Temperature sensors

Best Practice Advice on Cleaning:

Commonly used temperature sensors (thermistors) can be cleaned with “a detergent solution and a soft-

bristle brush” [5]. If recommended differently, the manufacturer’s guidelines on cleaning procedures must

be followed for both single and multi-probes.



Best Practice Advice on Calibration:

Several choices of calibrations can be explored depending on availability, facility and time. The temperature

sensors are quite durable and accurate and show low-drift characteristics. Temperature accuracy is especially

important because of the effect of temperature on the performance of other sensors. Calibrating

temperature sensors, a calibration thermometer (a non-mercury calibration thermometer can be used in

the field to check or monitor temperature readings) and a temperature-controlled water bath should be

used [5].

✓ The calibration thermometers should only be used for calibration rather than as field thermometers

[6].

An annual five-point calibration methodology is recommended over the temperature range of 0 to 40°C [5].

In addition, two-point calibration checks over the maximum and minimum expected annual temperature

range should be carried out at least every four months (three or more times per year) for thermistors [5].

✓ Quarterly or possibly monthly calibration can be required if the thermometer is in heavy use; exposed

to thermal shock, an extended period of direct sunlight, aggressive chemical solutions, or extreme

shifts in temperature.

A sample methodology for calibrating field thermometers with and without a commercial refrigerated water

bath is provided in Appendix VI-A: General Procedures for Calibration of Field Thermometers.

6.5.2.3 pH Sensor


In general, the only routine cleaning needed for (the body of) the pH electrode is rinsing thoroughly with

deionized water. However, in cases of extreme fouling (or contamination), the manufacturer’s instructions

on cleaning must be followed [5].


According to Wagner and colleagues (2006), “two standard buffer solutions bracketing the expected range

of environmental values are used to calibrate a pH electrode, and a third is used as a check for calibration

range and linearity of electrode response” [5]. Ionic buffer solutions (for pH 4, 7, and 10) are commonly used

in calibration of the pH instrument system. Buffers can resist changes to the specific pH value. For that

reason, the accuracy of the buffer solutions will have an impact on the accuracy of pH measurements.

✓ Use of buffer solutions with lower-than-standard molarity, in combination with the pH electrode with

a low ionic-strength, are recommended for dilute water with conductivity lower than 100 µS/cm [8].



✓ Use of buffer solutions with a higher-than-standard molarity is recommended for pH measurements

in high ionic-strength waters having conductivity greater than 20,000 µS/cm [8].

As a best practice advice, buffer bottles must be capped firmly after use to prevent evaporation and

contamination from atmospheric CO2.

✓ Sensitivity of standard buffers to CO2 contamination: pH 10 buffer > pH 7 buffer > pH 4 buffer.

✓ Variation of buffer pH with change in temperature: pH 10 buffer > pH 7 buffer > pH 4 buffer.

Prior to calibration, the temperature of the buffer solutions should be as close as possible to the mesocosm.

In order to ensure that, upon arrival at the field site, tightly capped buffer solutions should be immersed in

the mesocosms to allow time for temperature equilibration, usually 15 to 30 minutes.

✓ Waters with specific conductance values less than 100 µS/cm (low-ionic strength water) and greater

than 20,000 µS/cm (high-ionic strength water) require special buffers and pH sensors. The extra

preparations, precautions, and troubleshooting steps necessary to use these buffers and sensors to

measure low- or high-ionic strength waters are described in Ritz & Collins (2008) [8].

A sample calibration process for pH meters, including a wide range of available equipment, is compiled

from Wagner et.al. (2006) [5] and Ritz and Collins (2008) [8], which is provided in Appendix VI-B: General

Procedures for Calibration of pH Sensors of this SOP.

6.5.2.4 Optical DO Sensors

DO sensors are accessible from various manufacturers. As there are varieties DO sensors among

manufacturers, which are different in the instrument design and instructions for use, calibration and

maintenance, it is essential that one shall read the manual provided by the manufacturer thoroughly in

addition to the guidance provided in this SOP. Manufacturer’s guidelines must be followed, if these

recommendations deviate from the SOP.


Silt, outside the sensor, can be removed with a soft-bristle brush and the membrane should be wiped with a

lint-free cotton swab. DO sensors, then, should be rinsed with de-ionized water [5].


✓ The DO sensors should be temperature compensated. Because of the potential influence of altitude

and temperature, the DO probe(s) should be calibrated at the field.



✓ It should be noted that luminescent (optical) DO sensors are mostly calibrated by the manufacturer,

and the manuals indicate that calibration may not be required for up to a year. When calibrated, the

user should follow the manufacturer’s guidance. Regardless of the manufacturer’s claims, the user

must verify the correct operation of the sensor in the local measurement environment. The standard

protocol for servicing should be used for luminescent-based DO sensors to quantify the effects of

fouling and calibration drift. Rounds and colleagues advise the users to make frequent calibration

checks and to recalibrate as frequently as required to meet the specific data-quality objectives [9].

Recalibration should not be necessary if calibration checks show the sensor to be in agreement with

the calibration criteria [5].

DO sensors must be calibrated to 100-percent DO saturation and checked with a zero DO solution to provide

“an indication of sensor-response linearity” [5].

Three steps to calibration, as compiled from Wagner and colleagues (2006) [5] and Rounds and colleagues

(2013) [9], are provided in Appendix VI-C: General Procedures for Calibration of DO Sensors of this SOP.

6.5.2.5 Conductivity Sensor


First, the manufacturer’s recommendations must be checked before using acid solution or solvents on

sensors. Radtke and others (2004) recommend cleaning specific conductance sensors thoroughly with

deionized water [10]. A detergent solution can be used to remove oily or chemical residues. In addition, a

solvent or hydrochloric acid (5%) can also be used for dipping the sensor to be cleaned.

✓ The sensors can be soaked in detergent solution for hours without damage, however, care should be

taken when cleaning with an acid solution. The sensor must never be in contact with acid solution

for more than a few minutes.

✓ Sensors made of carbon and stainless-steel can be cleaned with a soft brush, but platinum-coated

sensors must never be cleaned with a brush.


Calibration and operating procedures differ, depending on the instrument. The manufacturer’s instructions

should be checked before starting calibration. The general procedures, that apply to most of the instruments

used in conductivity measurements, described in Radtke and colleagues (2004) [10] and [5] are summarized

in Appendix VI-D: General Procedures for Calibration of Conductivity Sensors of this SOP.

✓ For a cup-type cell, calibration and measurement procedures described for the dip-type cell apply; the

only difference is that standards are poured directly into the cup-type cell.

✓ “When using a dip-type cell, do not let the cell rest on the bottom or sides of the measuring

container” [10, p. 7].



6.5.2.6 Underwater Light (PAR) Sensor


Underwater light sensors are prone to biofouling. Accordingly, they should be cleaned frequently for

accurate measurements. The user should check the manufacturer’s guidelines for the best cleaning material.

As a best practice advice, debris, dust and other organic deposits on the PAR sensor lens should be removed

with water or window cleaner. One must not use an abrasive cleaner on the lens.

The salt accumulated on the sensor lens (especially in marine mesocosm) should be dissolved with vinegar

and removed from the lens with the help of a soft cloth or cotton swabs.

● Recording of pre- and post-cleaning results will help define the rate of sensor fouling, and the optimal

time between cleanings” [24].


Most of the PAR sensors are already calibrated prior to deployment at the factory and most of them are

recalibrated by the manufacturers when needed. For calibration, please refer to the manufacturer’s

instructions or contact with the manufacturer.

Quantum sensors (ie. LI-COR; LI-192 Underwater quantum sensor), are usually calibrated using a standard

light source (i.e. working standard quartz halogen lamps), which have been calibrated against reference

standard lamps. It is advised that every two years, the sensors should be calibrated by the manufacturer.

● The absolute calibration specification for quantum sensors is ± 5% (typically ± 3%).

As a best practice advice, the calibration can be verified using the Clear Sky Calculator at

www.clearskycalculator.com [27], which is an online calculator that reports theoretical photosynthetic

photon flux (PPF) at any time of day at any location in the world on a cloudless day (it is most accurate at

noon in spring and summer with completely clear skies).

6.5.2.7 Turbidimeter


Turbidity sensors are vulnerable to fouling especially in mesocosms high in sediment, algae accumulation,

larvae growth or other biological and chemical elements. Errors or drifts might occur also due to outgassing

air bubbles. The mechanical cleaning devices, wiper and shutters (most of the time, equipped with the

sensor), can be used to remove or prevent any accumulation.

✓ The probable algae accumulation on the wiper pad could prevent complete cleaning. Again,

accumulation of inorganic or organic debris in shutters could prevent operation of the sensor

http://www.clearskycalculator.com/



efficiently. For that reason, the wiper pad or other cleaning device also should be inspected for wear,

and cleaned or replaced if necessary.

As a best practice advice, the optic lens of the turbidity meter should be cleaned carefully with alcohol using

a soft cloth (or as recommended by the manufacturer), rinsed three times with turbidity-free water, and

carefully dried.

✓ If the readings are unusually high or erratic during the sensor inspection, entrained air bubbles may

be present on the optic lens and must be removed.


Before starting calibration, be certain that the probe is cleaned and free of debris. Solid particles left will

contaminate the standards during the calibration process and cause either calibration errors and/or

inaccurate field data. It should also be noted that turbidity meters should be calibrated directly rather than

by comparison with another sensor. The verification of calibration should be done with the same or similar

technology, at least in terms of the light source and the detection angle.

Field inspection or calibration of the turbidity sensor is made by using approved calibration turbidity and

calibration verification solutions (calibrants) and by following the manufacturer’s calibration instructions.

● The use of standards other than those mentioned in the manufacturer’s guidelines will result

in calibration errors and inaccurate field readings.

Calibration of the turbidity sensor should be checked in three standard solutions (although some instruments

may be limited to calibration with only one or two standards) before any adjustments are made.

✓ If instrument calibration allows only a two-step process, two primary standard solutions covering the

expected range must be used for calibration and a third midpoint standard solution is used to check

for linearity. Similarly, if the instrument calibration requires only turbidity-free water and one

standard solution, another midpoint standard solution must be used to check for linearity.

Turbidity calibrants can be instrument specific. Be careful to check the manufacturer's instructions. Use of

calibrants with instruments for which they are not designed can introduce significant errors. The three types

of turbidity calibrants generally recommended are (as summarized in Wagner and colleagues (2006), [5]):

(1) reference turbidity solutions, which are calibrants that are synthesized reproducibly from traceable raw

materials by a skilled analyst. The reference standard is fresh user-prepared formazin.

(2) calibration turbidity solutions, that are used in calibration must be traceable and equivalent to the

reference turbidity calibrants. Acceptable calibration turbidity solutions include commercially prepared

formazin, stabilized formazin, and styrene divinylbenzene (SDVB) polymer standards.



✓ Formazin-based calibrants can be diluted by using a dilution formula; however, errors may be

introduced during the dilution process, thus reducing the accuracy of the standard solution.

✓ Formazin-based calibrants are temperature dependent, and accurate readings may be difficult to

obtain during field conditions. Anderson (2005) [15] suggests that the effect of thermal fluctuations

can be minimized by calibrating the instrument at room temperature in an office laboratory using a

reference or calibration turbidity solution. Instrument calibration can then be checked at the field

site by using a calibration verification calibrant.

(3) calibration verification solutions and solids may include, but are not limited to, calibration turbidity

solutions; however, calibration verification calibrants that are sealed or solid materials must not be used to

adjust instrument readings [15].

✓ Before placing the sensor in a calibration verification calibrant, the sensor must be cleaned, rinsed

three times with turbidity-free water, and carefully dried. Turbidity-free water is prepared as

described by Anderson (2005) [15] and Wagner and colleagues (2006) [5].

6.5.2.8 Depth Sensor


Calibration of the depth sensors should be done in the atmospheric zero pressure and with respect to the

local barometric pressure.

For the calibration of depth sensors in multi probes, the user manual of the multi probes as well as the

software provided by the manufacturer should be visited. For instance, YSI EXO2 multi probes are equipped

with a non-vented strain gauge for measuring the depth. In order to calibrate the depth, the calibration

software provided with the multi probe should be used.

✓ It is important to note that, for the calibration of depth sensors for both vented or non-vented, the sensor

should be in air and not immersed in any solution.

6.5.2.9 Fluorometers (Chl-a)


Chlorophyll fluorescence is vulnerable to variations due to the environment changes, biofouling and

instrument design [28]. In order to improve and evaluate the accuracy, reliability, and stability of a

fluorometer, the optical surface should be inspected frequently. If needed, the optical surface should be

cleaned with a non-abrasive, lint-free cloth. One should take care to prevent scratches and damage to the

sensing window.




Pure Chlorophyll, dyes, algae cultures, and field water samples can be used as the calibration standards. Pure

Chl-a or liquid dyes such as Fluorescein Sodium Salt, Rhodamine WT Red, Rhodamine B, and Basic Blue 3

offered by various suppliers, can be used to calibrate fluorometers in the laboratory by standard methods.

Solid fluorescent materials, as trialed by Earp and colleagues (2011) are recommended as reference

standards for field calibration [28].

● Zeng and Li (2015) reminded that the dyes as well as pure or extracted Chl-a, have different

fluorescence intensity from the in vivo chlorophyll cell in natural populations [20]. They

recommended; in order to improve the accuracy among various species, pure phytoplankton cultures

should be used as a calibration sample (as in [29] and in [30]). Lawrenz and Richardson (2011) also

recommended the use of “natural communities collected from the site of interest” for calibration

of in situ fluorometers [31].

1 or 2-point calibration is recommended by most of the manufacturers. The probe/sensor can be calibrated

according to the measurement unit used in monitoring (μg/L or RFU; relative fluorescence unit from 0 -

100%). The RFU method is recommended if grab samples are also used to post-calibrate in vivo chlorophyll

readings.

For both measurement units, 2 reference standards should be used in calibration:

(1) clear (deionized or distilled) water (either 0 μg/L or 0%RFU)

(2) the standard solution with a known chlorophyll content OR dye solutions (not recommended in field

use; See the note in Calibration standards)

✓ Follow the manufacturer’s manual for the recommended standard solution/s and their

predetermined chlorophyll content/ratio.

The general procedures, that apply to most of the instruments used in situ chlorophyll measurements are

summarized in Appendix VI-E: General Procedures for Calibration of in situ Fluorescent Sensors. However,

Zeng and Li (2015) reminded that “the general and manufacturer calibration procedures are too simple to

meet scientific requirements; furthermore, calibration must be verified regularly due to species and

environment variation within space and time and lamp and sensor performance degradation over time” [20].

For that reason, fluorometers should be calibrated with multistep, pre- or post-calibration procedures, and

through special methods according to different situations. For more information and steps to the

recommended calibration procedure, please visit Zeng and Li (2015) [20].



6.5.2.10 ISE probes for measuring nitrogen


ISE-probes are (commonly) equipped with an automatic cleaning system using pressurized air, which proved

to work reliably [32].

● It is recommended to carry out a visual check of the probe before any calibration is started – if necessary

the probe should be cleaned manually [32].


A grab sample or a standard of known concentration can be used in calibration. The calibration standards

need to have sufficient ion activity; TISAB-solutions (Total Ionic Strength Adjustment Buffer) can be used to

adjust the ionic strength of the calibration samples [32].

✓ Supplementary sensors like temperature or pH should be calibrated before the ISE is calibrated, so that

any errors of the automatic temperature or pH-compensation are corrected before the ISE-calibration is

started [32].

✓ Sample readings should be taken after sensors have fully stabilized. Calibrating in a continuously stirred

solution from 1 to 5 minutes has shown to improve sensor performance (YSI EXO2, 2018).

✓ For best performance sensors should be calibrated as close to the expected field conditions as possible

(YSI EXO2, 2018).

✓ In multiparameter sensors, remove the ISEs to avoid exposing them to conductivity standards, Zobell

solution, pH buffer, or any solution with significant conductivity, as exposure to such solutions will reduce

data quality and response of the sensors.

Ammonium or nitrate sensors can be calibrated to one, two or three points. If preferred, a single-point

calibration procedure can be followed. However, the concentration of the measurement-ion at time of the

calibration should be in the upper half of the concentration range at the measurement location. In case the

maximum concentration of the measurement-ion is below 5 mg/l a two-points calibration should be carried

out in order to consider non-linearities in the lower measurement range” [32]. The 3-points calibration

method assures maximum accuracy and best performance of ISE sensors.

For the two- and the multiple-points calibration the probe has to be removed from the mesocosm and put

into a bucket with a grab sample or with a standard of known concentration.

✓ Using a grab sample of the actual measurement location has the advantage that influences due to

disturbance-ions at the time of sampling are compensated automatically. By using standards, reference

measurements can be omitted. However, a single-point calibration in-situ has to follow any calibration

in standards in order to consider influences from disturbance-ions.



For the two-points calibration the calibration measurements shall be carried out at approximately 20% and

80% of the concentration range of the measurement-ion at the measurement location and an approximate

concentration ratio of the calibration samples of 1:10. A multiple-points calibration can be applied in case

the concentration range at the measurement location has a wide span. Especially in the lower concentration

range of ammonium (<5 mgNH4-N/l) non-linearities have to be considered by means of a sectional linear

calibration function [32].

A sample 3-points calibration procedure for ammonium and nitrate is provided in Appendix VI-F: General

procedures for 3-points calibration of ammonium and nitrate sensors of this SOP.

✓ Ammonium and nitrate standards are good growth media for a variety of organisms. This growth can

significantly reduce the nitrogen content of your standards, an effect that is particularly important

for the 1 mg/L solution. It is best to use new standards for each deployment, but if you decide to save

your solutions for reuse, we recommend refrigerated storage to minimize the growth of these

organisms [19].

6.5.2.11 Multi Probes

As a best practice advice, the multi-probes should be calibrated in the following order (Adopted from [1]):

1. Dissolved Oxygen

2. Temperature (if not properly calibrated at the factory)

✓ Note that parameters such as pH, conductivity and oxygen are temperature sensitive and

therefore, it is important that the temperature being well calibrated.

3. Conductivity

4. pH

5. Depth

6. Light (PAR)

7. Chl-a (fluorescence)

8. Other parameters, if any

The calibration of each sensor should be carried out according to the procedure given in this SOP as well as

the guidelines provided in manufacturer’s handbook.



✓ When cleaning or calibrating a multi-probe, ensure access to a temperature stable and protected

location. Some chemical standards used to calibrate the multi-probe are temperature-sensitive and

it is always important to reduce the likelihood of contamination. In most cases, the data is collected

at the field site. Pre-cleaning data can be collected in mesocosms. The post-cleaning data can be

collected in water transported to the stable environment away from the mesocosm if possible, or

during redeployment (Adopted from [3]).

6.6 Overcoming Drift

6.6.1 Common Causes

Drift can be a major issue for deteriorating measurement quality. If the sensor is used in longer-term

deployments, drift is almost certain to occur. The extent of the drift will vary depending on the age of the

probe, the flow rate at the site, and the quality of the water as well as the quality of the instrument. The ion-

selective electrodes have the greatest tendency to exhibit calibration drift over time.

Optical DO sensors are stable and robust in maintaining calibration over long-term deployment and over a

wide range of environmental conditions. Accordingly, sensor (calibration) drift over time is minimal when the

sensor is kept clean.

Chlorophyll fluorescence (fluorometers) is also vulnerable to sensor drift and calibration rigor [28]. Errors or

drifts might occur in turbidimeters due to outgassing air bubbles.

Signal drift can be a major concern with on-line management systems because continuous immersion of the

sensors (especially ISEs) in water causes “electrode degradation, affecting the stability, repeatability, and

selectivity over time” [22].

✓ It should be noted that, some sensors may need regular manufacturer’s calibrations at a regular

interval. Otherwise, a user calibration may highlight a drift issue, in which case, sensors can be sent

back to the factory [24].

6.6.2 Elimination strategies

Sensor drift, in general, can be ameliorated by frequent calibration of the sensors manually. In addition, if

required, the authorized person should get in touch with the manufacturer as soon as possible.

To improve and evaluate the accuracy, reliability, and stability of a fluorometer and to eliminate the sensor

drift, the optical surface should be inspected periodically. In turbidimeters, the mechanical cleaning devices;

wiper and shutters (equipped with the sensor commonly) can be used to remove or prevent any

accumulation of air bubbles.



Sensor drift in the ion-selective electrodes can be ameliorated by using an ISE that has been conditioned in

the sample matrix, typically overnight, to produce a stable, reproducible and fast responding ISE, also

minimizing sample contact times and the concomitant ISE release of analyte into the sample” [33]. In

addition, the nitrate electrode drift can be fixed by “automatic calibration” [4]. For all monitoring studies

using ion-selective electrodes, the user should acquire a few grab samples during the deployment for analysis

in the laboratory or with another sensor that has been recently calibrated [19].

✓ Collecting grab samples from the site is encouraged to correct for drift [19].

6.7 Troubleshooting of sensors and record-keeping equipment

When a field parameter cannot be calibrated with specified methods, one shall determine the problem

source. If the problem is due to the sensor or the monitor, the necessary corrections should be made to

ensure the monitor is operational. For instance, spare sensors should be carried at every field visit so that

troubleshooting can be accomplished at the time of the visit. Accordingly, need for extra trips and record loss

can be prevented [5]. Some of the common problems that can be encountered in the field (and solutions)

are listed in Table 6-5.

Table 6-5: Common troubleshooting problems and likely solutions (adopted from [5])

Symptom Possible Problem Likely Solution

Temperature

Thermistor does not

read accurately

Dirty sensor Clean sensor

Erratic monitor

readings

Poor connections at monitor or sensor Tighten connections

Monitor slow to

stabilize

Dirty sensor Clean sensor

Readings off scale Failure in electronics Replace sensor or monitor

Dissolved oxygen

Meter drift or

excessive time for

monitor to stabilize

- Fouled sensor

- Wait for temperature equilibration

- Clean or recondition

- Check for obstructions or replace

Erratic monitor

readings

- Bad connection at monitor or

sensor

- Fouled sensor

- Tighten connections

- Clean or recondition

Monitor will not zero - Zero-DO solution contains oxygen

- Zero-DO solution is old

- Add additional sodium sulfite to zero-DO

solution

- Mix a fresh solution



Conductivity

Will not calibrate - Standard solutions may be old or

contaminated

- Electrodes dirty

- Air trapped around sensor

- Weak batteries

- Use fresh standard solutions

- Clean with soap solution

- Thrust sensors up and down and tap gently to

expel air

- Replace batteries

Erratic monitoring - Loose or defective connections - Tighten or replace connections

Monitor requires

frequent calibration

- Broken cables - Replace cables

- Replace monitors

pH

Meter will not

calibrate

- Buffers may be contaminated

- Faulty sensor

- Replace buffers

- Replace sensor

Slow response time - Dirty sensor bulb

- Water is cold or of low ionic

strength

- Clean sensor

- Be patient

Erratic readings - Loose or defective connections

- Defective sensor

- Tighten connections

- Replace sensor

Turbidity

Unusually high or

erratic readings

- Entrained air bubbles on the

optical sensor

- Damaged sensor

- Dirty sensor

- Water in connections

- Follow manufacturer’s directions

- Replace sensor

- Clean, following manufacturer’s directions

- Dry connector and reinstall

6.8 Quality Assurance and Quality Control (QA/QC) of high frequency measurement

Quality assurance during high-frequency measurements can be ensured by the following activities:

1. Timely and accurate documentation of field information in electronic and paper records: It is

recommended to keep a log book for each field instrument (if single probes are available) to record

the instrument repair, maintenance, and calibration history. The log book is recommended to be

made of a waterproof paper/material [34].

2. Use of manufacturer’s handbook, procedures and protocols to ensure sample integrity and data

quality: The manufacturer of the instrument(s) used for high frequency measurements should be

the primary source of information on the use, calibration, maintenance, troubleshooting and storage

of sensors. To ensure quality assurance and control, one shall read the manufacturer’s guidelines

thoroughly first. The recommendations provided in this section are general guidelines and best

practice advice related, prepared to ensure the level of quality assurance within the AQUACOSM

community.

3. Training of the personnel in charge in measurement techniques and the collection of quality-control



samples.

4. Second- or third-party auditing of such records, such as inter-(laboratory) calibrations.

Quality control during high-frequency measurements can be checked by the following activities:

1. Records of replicate measurements: In a QA/QC program for high frequency measurements,

determination of the “true value” of the measurement is fundamental [25]. The true value can be defined

by applying a second means of measuring with another instrument that is regularly serviced and

calibrated and only used for quality control purposes.

✓ Calibration standards may also be used as a reference value [25].

Other QA/QC applications at the field (adopted from [25])

“When real time or remote instruments are first placed into the field, accompanying measurements should

be taken with a handheld water quality instrument or by some other means. These field measurements –

which are sometimes paired samples taken for laboratory analyses – serve as QC points. These independent

field measurements (intermediate checks) are extremely important as they are the only check for the

accuracy and performance of the real time or remote water quality measurement. Each time an instrument

is removed and/or replaced, another complete set of field measurement should be collected” [25, p. 3].

Intermediate checks for a deployed multi-probe (eg., Hydrolab) should include (adopted from [1]):

1. “Upon deployment, field staff should collect a Winkler sample and, for other parameters, a check

measurement with another” instrument (if available, can be a hand-held instrument).

✓ “For DO, Winkler samples should be collected to bracket the expected high and low points for DO”.

2. For the other parameters, checks with another instrument” (if available, a hand-held meter) should be

conducted over the expected range of the parameter being measured.

✓ “At a minimum, one intermediate check should be completed for a short deployment (one week

or less)”.

3. The number of intermediate check measurements should be defined prior to the deployment and due to

the length of deployment [1].

4. At the end of deployment period, a final Winkler sample for DO should be collected and other check

measurements should be completed with an equivalent hand-held meter.

5. “This field check regime will provide a minimum of three checks per deployment and help identify if

instrument drift occurs” [1].



6.9 References 6 – High Frequency Measurements

[1] P. Anderson, “Standard Operating Procedures for

Hydrolab® DataSonde®, MiniSonde®, and HL4

Multiprobes,” 2016.

[2] Hach Company, “Hydrolab DS5X, DS5, and MS5

Water Quality Multiprobes,” 2006.

[3] Canadian Council of Ministers of the Environment,

“Protocols Manual for Water Quality Sampling in

Canada,” 2011.

[4] M. Beaupre, “Characterization of On-line Sensors for

Water Quality Monitoring and Process Control,”

Universite Laval Quebec, 2010.

[5] R. J. Wagner, R. W. J. Boulger, C. J. Oblinger, and B.

A. Smith, “Guidelines and standard procedures for

continuous water-quality monitors: Station

operation, record computation, and data reporting,”

2006.

[6] F. D. Wilde, “Temperature, (ver. 2.0),” 2006.

[7] B. Mostajir et al., “Use of sensors in marine

mesocosm experiments to study the effect of

environmental changes on planktonic food webs,” in

Sensors for ecology Towards integrated knowledge

of ecosystems, J.-F. Le Galliard, J.-M. Guarini, and F.

Gaill, Eds. Paris: CNRS, 2012, pp. 305–329.

[8] G. F. Ritz and J. A. Collins, “pH,” 2008.

[9] S. A. Rounds, F. D. Wilde, and G. F. Ritz, “Dissolved

Oxygen (Ver. 3.0),” in Geological Survey Techniques

of Water-Resources Investigations, Book 9, 3.0.,

Virginia: USGS, 2013, pp. 1-55.

[10] D. B. Radtke, J. V Davis, and F. D. Wilde, “Specific

Electrical Conductance,” 2004.

[11] E. Lewis, “The practical salinity scale 1978 and its

antecedents,” IEEE J. Ocean. Eng., vol. 5, no. 1, pp. 3–

8, 1980.

[12] K. Lee, A. Hassan, C. Lee, and J. Bae, “Microstrip

Patch Sensor for Salinity Determination,” Sensors,

vol. 17, no. 12, p. 2941, Dec. 2017.

[13] C. R. Booth, “The design and evaluation of a

measurement system for photosynthetically active

quantum scalar irradiance1,” Limnol. Oceanogr., vol.

21, no. 2, pp. 326–336, Mar. 1976.

[14] M. H. Long, J. E. Rheuban, P. Berg, and J. C. Zieman,

“A comparison and correction of light intensity

loggers to photosynthetically active radiation

sensors,” Limnol. Oceanogr. Methods, vol. 10, no. 6,

pp. 416–424, Jun. 2012.

[15] C. W. Anderson, “Turbidity,” 2005.

[16] E. Lannergård, “Potential for using high frequency

turbidity as a proxy for total phosphorus in Sävjaån,”

Swedish University of Agricultural Sciences

Department of Aquatic sciences and Assessment,

Uppsala, 2016.

[17] E. M. Ruzycki, R. P. Axler, G. E. Host, J. R. Henneck,

and N. R. Will, “Estimating sediment and nutrient

loads in four western lake superior streams,” JAWRA

J. Am. Water Resour. Assoc., vol. 50, no. 5, pp. 1138–

1154, 2014.

[18] M. Sadar, “Turbidity Measurement: A Simple,

Effective Indicator of Water Quality Change,” 2017.

[19] YSI, “EXO User Manual: Advanced Water Quality

Monitoring Platform,” 2018.

[20] L. Zeng and D. Li, “Development of In Situ Sensors for

Chlorophyll Concentration Measurement,” J.

Sensors, vol. 2015, pp. 1–16, 2015.

[21] M. Babin, “Phytoplankton fluorescence: theory,

current literature and in situ measurement,” Real-

Time Coast. Obs. Syst. Mar. Ecosyst. Dyn. Harmful

Algal Bloom., pp. 237–280, 2008.

[22] W. J. Cho, D.-W. Kim, D. H. Jung, S. S. Cho, and H.-J.

Kim, “An Automated Water Nitrate Monitoring

System based on Ion-Selective Electrodes,” J.

Biosyst. Eng., vol. 41, no. 2, pp. 75–84, Jun. 2016.

[23] C. Borden and D. Roy, “Water Quality Monitoring

System Design,” Manitoba Canada, 2015.

[24] E. de Eyto, M. Dillane, A. Laas, D. Pierson, and E.

Jennings, “Station maintenance (Factsheet 012),” in

NETLAKE Guidelines for automatic monitoring

station development. Technical report. NETLAKE

COST Action ES1201, E. Laas, A., de Eyto, E., Pierson,

D. and Jennings, Ed. 2016, pp. 47–49.

[25] OTT Hydromet, “Quality Assurance and Quality



Control: Technical Note,” 2017.

[26] Ocean Optics, “How accurate is a spectrometer

wavelength calibration light source from Ocean

Optics?,” 2019. [Online]. Available:

https://oceanoptics.com/faq/calibration-light-

source-accuracy/. [Accessed: 05-May-2019].

[27] Apogee Instruments, “Clear Sky Calculator | Apogee

Instruments Inc.” [Online]. Available:

http://www.clearskycalculator.com/. [Accessed: 04-

May-2019].

[28] A. Earp et al., “Review of fluorescent standards for

calibration of in situ fluorometers:

Recommendations applied in coastal and ocean

observing programs,” Opt. Express, vol. 19, no. 27, p.

26768, Dec. 2011.

[29] E. J. D’Sa, S. E. Lohrenz, V. L. Asper, and R. A. Walters,

“Time Series Measurements of Chlorophyll

Fluorescence in the Oceanic Bottom Boundary Layer

with a Multisensor Fiber-Optic Fluorometer,” J.

Atmos. Ocean. Technol., vol. 14, no. 4, pp. 889–896,

Aug. 1997.

[30] C. W. Proctor and C. S. Roesler, “New insights on

obtaining phytoplankton concentration and

composition from in situ multispectral Chlorophyll

fluorescence,” Limnol. Oceanogr. Methods, vol. 8,

no. 12, pp. 695–708, Dec. 2010.

[31] E. Lawrenz and T. L. Richardson, “How Does the

Species Used for Calibration Affect Chlorophyll a

Measurements by In Situ Fluorometry?,” Estuaries

and Coasts, vol. 34. Coastal and Estuarine Research

Federation, pp. 872–883, 2011.

[32] S. Winkler, L. Rieger, E. Saracevic, A. Pressl, and G.

Gruber, “Application of ion-sensitive sensors in

water quality monitoring,” Water Sci. Technol., vol.

50, no. 11, pp. 105–114, 2004.

[33] R. De Marco, G. Clarke, and B. Pejcic, “Ion-Selective

Electrode Potentiometry in Environmental Analysis,”

Electroanalysis, vol. 19, no. 19–20, pp. 1987–2001,

Oct. 2007.

[34] F. D. Wilde, “Guidelines for field-measured water-

quality properties,” 2008.

Co-funded by the European Union D4.1 Standard Operating Procedures| 7 Data Quality Assurance and Quality Control


7. Data Quality Assurance and Quality Control

7.1 Definitions and terms

TERM DEFINITION(S)

Flags A system to identify the quality of data which preserves the original data and

indicates the degree of manipulation

Metadata Contextual information to describe,understand and use a set of data. [2]

QA & QC Quality Assurance and Quality Control is a two-stage process aiming to identify

and filter data in order to assure their utility and reliability for a given purpose

QA Quality Assurance (process oriented)

Is process oriented and encompasses a set of processes, procedures or tests

covering planning, implementation, documentation and assessment to ensure

the process generating the data meet a set of defined quality objectives.

QC Quality Control (product oriented)

Is product oriented and consists of technical activities to measure the attributes

and performance of a variable to assess whether it passes some pre-defined

criteria of quality.

7.2 Cross reference

All other SOP’s provided by AQUACOSM in which data are collected should refer to this SOP in relation to QA

procedures.

Materials and Reagents

● Software as Excel, R, SPSS, SAS, Systat or other statistical programs. QC procedures may also be built

into database functionality.

7.3 Health and safety regulation

Not relevant.



7.4 Environmental indications

Not relevant.

7.5 Quality Assurance and Quality control Workflow

The purpose of quality assurance and quality control is to ensure the reliability and validity of the information

content of the data. Many QA & QC measures can be undertaken; however, a ubiquitous and crucial

characteristic is that each step is documented, described and repeatable. Documentation is the key to make

data reliable, valuable [3] and reusable [4]. The figure below describes the recommended workflow for

AQUACOSM data collection.

Figure 7-1: Suggested QA and QC workflow

Based on the level of quality assurance and control steps the raw data has undergone, we distinguish four data levels:

level 0 - raw data

level 1 - automated QC - large obvious errors removed

level 2 - manual QC

level 3 - Gap filled or Interpolated data

level 4 - aggregated and summarized data



7.5.1 Quality assurance of raw data collection

The quality of the data to be collected in the mesocosm experiment will be improved when the data collection

preparation and mesocosm sampling undergoes several quality assurance steps. These quality assurance

steps differ for the type of data collected, and more detailed information on quality assurance for specific

sampling procedures can be found in the SOPs for manual data collection (zooplankton, water chemistry,

phytoplankton, periphyton, protozoa, bacteria, archaea, viruses etc). See above for examples, including

sensor calibration, and adequate labelling of sampling containers.

In addition to method-specific quality assurance steps as described in the specific SOP s, prior to collecting

the data, one should define standard names for common objects. This should be done for at least the

following cases:

● Parameter name: use standardized names that describe the content and describe the parameter in

the metadata [1, 3, 5]. As a best practice advice: use the vocabulary developed during the

AQUACOSM project with the standardized names.

● Formats: choose a format for each parameter, describe this format in the metadata and use it

through the whole dataset. Important formats to consider are dates, times, spatial coordinates and

significant digits. [1, 5]

● Taxonomic nomenclature. Follow international species data list.

● Measurement units: make use of the SI units (and the AQUACOSM vocabulary) and document these

units (in the metadata). [1, 5]

● Codes: “standardized list of predefined values”. Determine which codes to use, describe the codes

and use the codes consistently. Every change that is made in the codes, should be documented. [1,

5]

● Metadata: data about data, with as goal to help scientists to understand and use the data [1]. The

mesocosm metadatabase developed in Aquacosm (link) is currently build on the Ecological Metadata

Language (EML, see Fegraus et al. 2005), specifically adapted to mesocosm data.

Another important point of QA is to assign the responsibility for data quality to a person or persons who has

some experience with QA & QC procedures [1].

7.5.2 Quality Control

Raw or primary data should not be removed or changed unless there is solid evidence that it is erroneous. In

the first instance questionable data should be flagged according to international code system (e.g. ICES or

the like). In the event that primary data are altered it must be saved and a motivation for the action added

in the same post. It is essential that the raw, unmanipulated form of the data is saved so that any subsequent

procedures performed on the data can be repeated [6]. Instead of removing or deleting data it is preferable

to use a system of flags, via a range of QA processes and steps, thereafter QC can be carried out by filtering

the data based on the flags and further analysis carried out.

https://www.aquacosm.eu/geonetwork/srv/eng/catalog.search#/home



The general QC checks that should be done are described in detail in [7], this can be done manually or

automated, the latter being more applicable to high frequency data:

● Gap or missing value check: do you have all the expected results?

● Control samples: Are they within expected range and variability?

● Calibration curves: Are coefficients within expected range and variability?

● Spurious / impossible results check: do you have negative results or results with an unreasonable

magnitude?

● Outlier check: do the results fall outside an expected distribution of the data?

● Range check: do the calculated values fall within the expected range and variability?

● Climatology check: are the results reasonable compared with historical results (range and patterns)?

● Neighbour check: are the results reasonable compared to results of the same site, same day or

different depths?

● Seasonality check: do results reflect seasonal processes or effects or are they extraordinarily

different?

For a sensor network QC should include [6]:

● Date and time: check if each data point has the right date and time.

● Range: check if data fall within established upper and lower bounds

● Persistence: check if the same value is recorded repeatedly, this can indicate problems like a sensor

error or system failure.

● Change in slope: check the change in slope to see if the rate of change is realistic for the type of data

collected

● Internal consistency: evaluate differences between related parameters

● Spatial consistency: check replicate sensors or compare the sensors with identical sensors from

another site.

After the QC check one should evaluate data points that did not pass the QC check. Compare with other

variables, check calibration curves and other types of sampling and instrument performance. Check log books

for comments by the responsible person. Only when evidence for sampling error, contamination or

instrument failure is shown should data point be removed or replaced with a, for example interpolated value.

The decision and its motivation must be documented and the data point flagged accordingly.

7.5.2.1 Flagging

This SOP states that to establish successful QC procedures it is crucial to flag your data to explain differences

between the raw data and processed data. “Flags or qualifiers convey information about individual data

values, typically using codes that are stored in a separate field to correspond with each value” [6].

We recommend using the flagging values as suggested by Hook et al. [5], as it allows you to retrieve the QC

steps your data has undergone.



Comment: other possible flagging systems are: Campbell et al. [6], Marine water website [8], QARTOD

flagging system [9], IODE flagging system by Reiner Schlitzer [10] and protocols produced EU infrastructure

projects the Copernicus Marine Environment Monitoring Service (CMEMS) [11]. Table 7-1 suggests a flagging

system that may be useful for mesocosm applications. It is more complicated than some systems but it

contained information of why a data point is questionable and allows filtering of the data with different

stringency of quality control.

Table 7-1: Recommend flag values [5] for the AQUACOSM project

Flag Value Description

V0 Valid value

V1 Valid value but comprised wholly or partially of below detection limit data

V2 Valid estimated value

V3 Valid interpolated value

V4 Valid value despite failing to meet some QC or statistical criteria

V5 In-valid value but flagged because of possible contamination (e.g., pollution source,

laboratory contamination source)

V6 In-valid value due to non-standard sampling conditions (e.g., instrument malfunction,

sample handling)

V7 Valid value but set equal to the detection limit (DL) because the measured value was

below the DL

M1 Missing value because not measured

M2 Missing value because invalidated by data originator

H1 Historical data that have not been assessed or validated

Table 7-2: The ARGO data quality flagging system


0 No data quality control on data

1 Data passed all tests

2 Data probably good

3 Data probably bad. Failed minor tests

4 Data bad. Failed major tests

7 Averaged value

8 Interpolated value

9 Missing data



Table 7-3: ICES Data Quality Flag


0 data are not checked

1 data are checked and appear correct

2 data are checked and appear inconsistent but correct

3 data are checked and appear doubtful

4 data are checked and appear to be wrong

5 data are checked and the value has been altered

7.5.2.2 Automated QC

The large increase in the number of studies using multiple sensors recording at high frequency has led to the

generation of large volumes of data, making it extremely difficult and undesirable to carry out manual QC. A

number of automated methods have been developed for quality control. Visual checks of such data are

relatively straightforward, if time consuming. Therefore, it is increasingly desirable to automate these

approaches, which ideally would be integrated into database functionality, for example through the use of R

or Python code, adapted to the particular data set. These QC steps can be applied in real time or post data

collection. Real time quality control (RTQC) has the advantage that an alarm system can be integrated to

indicate suspect values providing an early warning of sub-optimal sensor performance.

These methods follow many of the same QC methods as manual methods, in particular:

● Date and time: this checks if each data point has the right date and time.

● Range: these check if data fall within established upper and lower bounds can be divided into Global

range tests (possible distribution) and local range tests (probable distribution)

● Persistence: or frozen value test - this checks if the same value is recorded repeatedly, this can

indicate problems like a sensor error or system failure.

● Change in slope or spike test: this checks the change in slope to see if the rate of change is realistic

for the type of data collected

● Internal consistency: this evaluates differences between related parameters

● Spatial consistency: this checks replicate sensors or compare the sensors with identical sensors from

another site.

Parameters measured by autonomous sensors can vary on a range of scales and a number of sensor types,

in particular optical sensors, can be affected by non-negligible noise. There are a number of methods which

can be used to identify outliers or spurious data that can then be flagged accordingly and filtered out of

subsequent analysis. One approach [11] has been to apply a procedure testing the statistical entropy caused

by each progressive measurement, as described by [12], this is a 2-step estimation of the Akaike information

criterion - details can be found in [12]. This approach to outlier or spike identification is highly dependent on



the number of sample points considered in the estimate of statistical entropy. The inclusion of too few

samples risks the exclusion becoming too sensitive with ‘good’ data being excluded. The inclusion of too

many samples and the test may potentially become too insensitive (Figure 7-2). In addition, the designation

of an observation as an outlier is also dependent on the selection of a cut off or critical value of variation

above which a value is designated as an outlier. The selection of these two parameters for the test is

dependent upon the type of data being collecting. For example, the data generated by the Ferrybox [11] are

collected 400 m apart and in this case, it was necessary to relax the criteria used for identifying outliers as

their natural variation in samples 400 m apart is greater than samples measured in the same location.

A similar approach to identification of outliers in sensor data has been developed at the University of

Waikato, New Zealand (https://www.lernz.co.nz/tools-and-resources/b3) and now also available as an R

package (https://github.com/kohjim/rB3) , under the aegis of the GLEON network. This is a freely available,

downloadable programme that can be used for post-data collection data processing. B3 is an integrated

programme that can be used to carry out many of the QC steps outlined above, e.g. range check, missing

data, repeated values. It also identifies outliers or spikes samples by two methods. The first is similar to the

above described method, where a running mean is calculated and observations that fall outside a critical

standard deviation can be identified as potential outliers. Both the period over which the running mean is

calculated and the cut off value of the standard deviation can be altered to suit the dataset in question. The

second method uses a rate of change analysis (ROC) to identify jumps in the data that are out of the normal

range. The number of data points used to identify the ‘normal’ range can be altered as can the critical rate

of change value above which a value is deemed a potential outlier. Each of these methods has different

sensitivities to outliers, depending on the kind of data analysed and there is a balance to strike between

excluding ‘good’ data and including ‘bad’ data. For example, data with a large diurnal range (e.g. DO data)

may be vulnerable to excluding good data at the high and low end of the diurnal cycle when the running

mean methods are used. Thus, it may be necessary to tailor the cut off values for ROC analysis and running

mean analysis for each dataset, or even each mesocosm and some form of manual QC is highly

recommended.

https://www.lernz.co.nz/tools-and-resources/b3

https://github.com/kohjim/rB3



Figure 7-2: Ferry box turbidity data with spike point analysis described in section 7.2.2 from [11,12] showing the effects of including different numbers of observations (5, 10 and 100). The red points are identified as outliers and n=5 and n=10 appear too sensitive and exclude a large amount of good data.



7.5.3 Aggregated, summarized data

As a final step, data can be aggregated and summarized for publication purposes. This includes integration

over e.g. depth (space) and time. It should be remembered that integration however reduce the degrees of

freedom and may limit the statistical tests that can be performed and possibly the statistical power.

Aggregation is preferably done in a database environment with the integrating function located at one

instance. This function should be validated by manual calculation with selected data from the same set.

The start and end values for the range of integration should be clearly defined, as is true for methods for

inter- and extrapolation where applicable.

Number of samples included in the integrated value (n) and its standard deviation (±SD) can be provided to

assess the extent of data for the aggregated value.



7.6 References 7 – QA &QC

1. Michener, W.K. and M.B. Jones, Ecoinformatics:

supporting ecology as a data-intensive science. Trends in

Ecology & Evolution, 2012. 27(2): p. 85-93.

2. Jones, M.B., et al., The New Bioinformatics:

Integrating Ecological Data from the Gene to the Biosphere.

Annual Review of Ecology, Evolution, and Systematics, 2006.

37(1): p. 519-544.

3. Rüegg, J., et al., Completing the data life cycle: using

information management in macrosystems ecology research.

Frontiers in Ecology and the Environment, 2014. 12(1): p. 24-

30.

4. Michener, W.K., Ecological data sharing. Ecological

Informatics, 2015. 29: p. 33-44.

5. Hook, L.A., et al., Best practices for preparing

environmental data sets to share and archive. 2010, Oak

Ridge National Laboratory Distributed Active Archive Center,

Oak Ridge, Tennessee, U.S.A. p. 40.

6. Campbell, J.L., et al., Quantity is Nothing without

Quality: Automated QA/QC for Streaming Environmental

Sensor Data. BioScience, 2013. 63(7): p. 574-585.

7. Bos, J., C. Krembs, and W.R. Kammin, EAP088

Marine Waters Data Quality Assurance and Quality Control V

1.0 5/30/2015. 2015: p. 35.

8. Department of Ecology - State of Washington.

Marine Waters Data Quality Codes. [webpage] 2017 [cited

2017 October 17]; Available from:

http://www.ecy.wa.gov/programs/eap/mar_wat/datacodes.

html.

9. Integrated Ocean Observing System. Manual for

Real-Time Oceanographic Data Quality Control Flags. 2017

[cited 2017 October]; Available from:

https://ioos.noaa.gov/wp-

content/uploads/2017/06/QARTOD-Data-Flags-

Manual_Final_version1.1.pdf.

10. Schlitzer, R. Oceanographic quality flag schemes

and mappings between them. 2013 2013-05-24 [cited 2017

October]; version 1.4:[Available from:

https://odv.awi.de/fileadmin/user_upload/odv/misc/ODV4_

QualityFlagSets.pdf.

11. Jaccard Pierre, Hjemann Dag Oystein, Ruohola Jani,

Ledang Anna Birgitta, Marty Sabine, Kristiansen Trond, Kaitala

Seppo, Mangin Antoine (2018). Quality Control of

Biogeochemical Measurements. CMEMS-INS-BGC-QC.

https://doi.org/10.13155/36232

12. Ueda, T. 2009. A simple method for the detection of

outliers. Electronic Journal of Applied Statistical Analysis 2:67-

76.

http://www.ecy.wa.gov/programs/eap/mar_wat/datacodes.html

http://www.ecy.wa.gov/programs/eap/mar_wat/datacodes.html

https://ioos.noaa.gov/wp-content/uploads/2017/06/QARTOD-Data-Flags-Manual_Final_version1.1.pdf



https://odv.awi.de/fileadmin/user_upload/odv/misc/ODV4_QualityFlagSets.pdf

https://odv.awi.de/fileadmin/user_upload/odv/misc/ODV4_QualityFlagSets.pdf

https://doi.org/10.13155/36232

https://doi.org/10.13155/36232

https://doi.org/10.13155/36232

Co-funded by the European Union D4.1 Standard Operating Procedures| III Appendix


III. Appendix

Appendix I: Phytoplankton

The standard recipe for Lugol’s Solution (Adopted from [46])

Equipment and Supplies

✓ Reagent water

✓ Glacial acetic acid

✓ Iodine

✓ Potassium iodide (KI)

✓ 1 L Volumetric flask

✓ Opaque 1 L container

✓ Glass funnel

Lugol’s Solution preparation procedure

Step 1: In a fume hood, dissolve, 100 g of KI and 50 g of I2 in approximately 800 mL of reagent water in a 1 L

volumetric flask.

Step 2: Mix until the chemicals are completely dissolved.

Step 3: Add 100 mL of glacial acetic acid and bring volume up to 1 L with reagent water. Add 1 mL of Lugol’s

solution to each 100ml water sample within 2 hours of sample collection to obtain a final concentration of

1%. Store the solution in an opaque or brown glass container.

Important considerations

● Lugol’s solution should be prepared maximum one week prior to the survey.

● Lugol’s solution is stored in an opaque bottle labeled with the contents, date of preparation, and

preparer’s initials.



Recipes for alkaline, neutral and acidic Lugol’s solution from [16]

Formaldehyde Stock solution (Adopted from [46])

Equipment and supplies

✓ Normal formaldehyde solution 40% v/v

✓ Deionized water

✓ 1 L Volumetric flask

✓ 1 L container

✓ Glass funnel

Formaldehyde Stock Solution preparation procedure

Step 1: Acquire the normal formaldehyde solution 40% v/v

Step 2: In a fume hood, measure 400 mL of the normal formaldehyde solution with a volumetric cylinder and

add it in a 1 L volumetric flask using a glass funnel

Step 3: Fill the flask with water to obtain 1 L of Formaldehyde stock solution

Step 4: Transfer the contents of the flask in a Plastic container and cap it firmly. Store under a fume hood.

Important considerations

● Formaldehyde is carcinogenic and should be handled with care. Proper safety equipment includes

gloves and plastic goggles.

● The preparation of the formaldehyde Stock Solution should be done in a fume hood.



Appendix I-B

Label Standard (Revised from [46])

Mesocosm Facility: ………………………………......... Mesocosm Number: …………………………………

Sample No:………………………………………………...... Sampler: ……………………………………………………

Secchi Depth: ………………………………………………. Max. Sampling Depth: ………………………………..

Sampling Depths: ………………………………………………………………………………………………………………..

Date and Time: …………………………………………….. Collector Initials: ……………………………………….



Phytoplankton Sampling Field Datasheet (Revised from [46])

Phytoplankton Sampling Field Sheet

Mesocosm Name: ……………………………………………………………………………………………………………….

Mesocosm Number……………………………………… Date and Time: ……………………………………………

NOTES:

NAME & SIGNATURE:



Appendix II: Zooplankton

Appendix II-A

Table A – 1. The advantages and disadvantages of preservatives (Tabularized from [1] )

Preservative

solution

Advantages Disadvantages

Lugol’s

Iodine

● It enters the cell more quickly than

formaldehyde, leaving shock-sensitive

organisms better preserved in the

sample.

● Detection of small mesozooplankton is

easier, thanks to the enhanced

contrast between organisms and the

surrounding fluid.

● Slightly toxic in normal conditions of

use and offers minimal discomfort to

laboratory personnel.

● The intense staining may obscure certain cellular structures

that need to be observed for proper identification (e.g.

surface structures). Overstaining can be cleared up by adding

sodium thiosulphate, which reduces the iodine.

● Organic matter is not fixed, and soft materials may lose their

characteristic structure during storage of the sample. This

alteration can be prevented by the addition of

formaldehyde.

● Iodine is oxidised over a period of time; therefore, when

storage time is long or samples contain a large amount of

organic matter, samples need attention to prevent them

from decay.

Formaldehyd

e (Formalin)

● Effective prevention of the microbial

degradation of organic matter.

● Organic structures and other

morphological characteristics remain

visible.

● When stored properly in suitable

bottles, samples will stay in good

condition for many years without

attention.

● Distortion of the body structure takes place especially in soft-

bodied organisms (e.g. several rotifers and cladocerans)

● Cladocerans (e.g. Daphnia, Bosmina. Diaphanosoma) may

balloon followed by the loss of brood-pouch contents (eggs

and embryos). This can be prevented by using a cold (6 °C)

solution of formalin with added sucrose.

o In this case, it is advisable to kill the organisms using more

rapid and efficient methods or to utilize special solutions and

techniques [18].

● Formaldehyde is irritating at even very low concentrations.

Formaldehyde can also be carcinogenic. This preserving

solution should therefore be handled with care, and should

be “washed out” of samples within an air extraction hood

before they are analysed.

Ethanol ● The quality of the samples can be

retained for long periods of time if

they have been stored correctly.

● Use of 96% ethanol prevents carapace

distortion and loss of eggs and

embryos due to ballooning in

cladocerans

● DNA is retained in a form which can

subsequently be extracted for genetic

analysis.

● Cell shrinkage can be observed, which will result in

underestimation of dimensions.

● May be unpleasant for laboratory personnel (dizziness,

headaches). For this reason, this preserving agent should be

diluted before samples are analysed.



How to prepare the reagents?

a) Lugol’s solution (Adopted from [1] )

✓ Distilled/demineralized water

✓ Potassium iodide (KI)

✓ Iodine (crystalline)

✓ Glacial acetic acid

Lugol’s Solution preparation procedure

Dissolve 100 g KI (potassium iodide) in 1 l of distilled or demineralized water; then add 50 g iodine

(crystalline), shake until it is dissolved and add 100 ml of glacial acetic acid. As this solution is close to

saturation, any precipitate should be removed by decanting the solution before use.

b) Formaldehyde Solution Preparation Procedure (Adopted from [1] )

Formaldehyde is neutralized, e.g. with hexamethylenetetramine (C6H12N4). Dilute the formaldehyde with

water to 20 % (v/v) to avoid precipitation, and then add 100 g of hexamethylenetetramine and 40 g to 80 g

sucrose per litre of 20 % formaldehyde.

c) Buffered sucrose formation (Buffered Formalin) (Adopted from [1] and [5]):

✓ Sucrose (crystalline),

✓ Formalin (37% solution of formaldehyde in water),

✓ Borax (powder)

Buffered Formalin preparation procedure

Dissolve 60 grams of sucrose in 1000 mL of formalin [97]. Then, dissolve 9 grams of borax in 1000 mL of

sucrose formalin. Store in a labelled plastic container.



Appendix II-B



Appendix II-C

Counting cladocerans

✓ Open the excel file “Counting_cladocerans” on the computer and make sure the macro is enabled.

✓ Take some time to get acquainted with the file.

Note: On “Sheet 1” there’s a row with 50 cladoceran species, a column for the input of sample codes

and 14 buttons. These buttons are all working as soon as the macro is enabled, but to activate

or deactivate a quick key function of the buttons with the numbers 1-9 and “num enter” you

can use Control+Q or Control+W respectively, or you can click on the buttons “Activate Quick

Keys” or “Deactivate Quick Keys”. Once activated the buttons 1-9 correspond to the keys 1-9

on the keyboard and the numpad of the computer and the button “num enter” corresponds

to the enter key on the numpad. The buttons can be dragged and dropped to a different

location by right clicking to select them and then left clicking to drag and drop them.

✓ Pressing a button will enter +1 to the cell corresponding to that button and will also enter data in

“Sheet 2” (more information on “Sheet 2” follows). Which cell corresponds to the button depends on

the location of the button and the location of a preselected cell. The column the button resides in is

the column the data will end up in and the preselected cell determines the row of the cell the data

will end up in (this can be any cell in the row).

✓ Each time a button is pressed in “Sheet 1” “Sheet 2” counts the number of species and individuals in

the selected row and stores that information in two columns, aptly named “COUNT_SPECIES” and

“SUM_INDIVIDUALS”. These two data points are then added to a graph right next to the two

aforementioned columns, thereby creating a saturation curve.

✓ “Sheet 3” named “ml” has columns for e.g.: recording the starting volume of and the amount of the

subsample taken from the sample and “Sheet 4” named “All_Saturation_Curves” is there for

transferring the two columns of “Sheet 2” to, after a sample is finished.

✓ Activate the quick keys and start counting the cladocerans. Drag and drop the buttons 1-9 and “num

enter” to the columns of the most encountered species for optimal speed.

✓ Use the two dissection needles to turn and if necessary, to dissect the cladocerans.

✓ When the identification of a certain individual with the stereomicroscope is impossible, use the glass

Pasteur pipette to transfer the individual to the microscope depression slide. Add a drop of water to

fill the depression and use the tissue to remove access water after covering the depression with the

cover slip.

✓ After identification transfer the individual from the slide to the small third Petri dish.

✓ When all the individuals in the subsample have been counted, transfer the contents of the gridded

(and the small, if the subsample counted was the last one) Petri dish to a clean culture tube labeled

“sample_name done”.

Note: check with the stereomicroscope if all mesozooplankton was transferred, often some get stuck

to the bottom of the Petri dish.



✓ Continue taking subsamples and counting until the saturation curve (see “Sheet 2” of the excel file

“Counting_cladocerans”) levels out. Since levelling out is a somewhat flexible concept, counting 100

individuals without finding a new species is hereby set as the definition of this concept. This means

counting can be stopped as soon as a whole subsample has been counted and no new species were

found for the last 100 individuals.

✓ When starting a new sample make sure that all materials are clean of mesozooplankton by

thoroughly rinsing them.



Appendix II-D

Key literature to identify Crustacea:

● L. A. Błędzki and J. I. Rybak, “Freshwater crustacean mesozooplankton of Europe,” Switz. Springer,

2016. [98]

● L. M. Witty, “Practical guide to identifying freshwater crustacean mesozooplankton.” Cooperative

Freshwater Ecology Unit, 2004. ([99])

● U. Einsle, Guides to the Identification of the Microinvertebrates of the Continental Waters of the

World. 1996. ([100])

Key literature to identify Rotifer:

Identification of rotifers should proceed with the use of taxonomic keys. Taxonomic identification, commonly

followed are as follows:

● E. McCauley, “The estimation of the abundance and biomass of mesozooplankton in samples,” in A

manual on methods for the assessment of secondary productivity in fresh waters, 1984, pp. 228–265.

([49])

● A. Ruttner-Kolisko, “Suggestions for biomass calculations of planktonic rotifers,” Arch. fur Hydrobiol.

Beihefte, vol. 21, pp. 71–76, 1977. ([76]) (species)

● K. W, Rotatoria. Die Radertiere Mitteleuropas Gebruder Borntraegerand. Berlin, 1978. ([101])

(species)

● W. T. Edmondson, “Rotifera,” Freshw. Biol., pp. 420–494, 1959. ([102])

● R. S. Stemberger, A guide to rotifers of the Laurentian Great Lakes, vol. 1. Environmental Monitoring

and Support Laboratory, Office of Research and Development, US Environmental Protection Agency,

1979. ([103]). (genera)

● R. W. Pennak, “Fresh-water invertebrates of the United States 3rd Edition,” in Fresh-water

invertebrates of the United States, 3rd ed., New York: Wiley New York, 1989. ([104])

● R. L. Wallace, “Rotifera,” eLS, 2001. ([105]) (freshwater Rotifera)

Other taxonomic identification literature

● H. Bottrell et al., “A review of some problems in mesozooplankton production studies,” Nor. J. Zool.,

vol. 24, pp. 419–456, 1976. ([48])

● Mekong River Commission and Environment Programme, “Identification Handbook of Freshwater

Zooplankton of the Mekong River and its Tributaries Identification Handbook of Freshwater

Zooplankton of the Mekong River and its Tributaries,” 2015. ([106])

● J. H. Thorp and A. P. Covich, Ecology and classification of North American freshwater invertebrates.

Academic press, 2009. ([107])

● T. Nogrady, R. Pourriot, and H. Segers, “Rotifera 3. Notommatidae and Scaridiidae,” Guid. to Identif.

Microinvertebrates Cont. Waters World 8.(H. Dumont, T. Nogrady, eds). SPB Acad. Publ. BV, 248 p.,

1995. ([108]).



● H. Segers, “Rotifera 2. The Lecanidae (Monogononta),” Guid. to Identif. Microinvertebrates Cont.

Waters World 6.(HJ Dumont, T. Nogrady, eds). SPB Acad. Publ. BV., 226 p., 1995. ([109])

● W. De Smet, “The Proalidae (Monogonanta),” in Guides to the identification of the

macroinvertebrates of the continental waters of the world, H. Dumant, Ed. Amsterdam: SPB

Academic publishing, 1995, pp. 1–102. ([110])

● W. De Smet and R. Pourriot, “The Dicranophoridae (Monogononta) and Ituridae (Monogononta),” in

Rotifera Vol. 5, Guides to the identification of the macroinvertebrates of the continental waters of the

world, H. Dumant, Ed. The Hague: SPB Academic publishing, 1997, p. 1–344. ([111])

● K. Smith and C. H. Fernando, A guide to the freshwater calanoid and cyclopoid copepod Crustacea of

Ontario. Department of Biology, University of Waterloo, 1978. ([112])

● M. D. Balcer, N. L. Korda, and S. I. Dodson, Zooplankton of the Great Lakes: a guide to the

identification and ecology of the common crustacean species. Univ of Wisconsin Press, 1984. ([113])

● J. L. Brooks, “The systematics of North American Daphnia,” Mem. Connect. Acad. Art Sci., vol. 13, pp.

1–180, 1957. ([114])

● B. Dussart, Les Copépodes des eaux continentales d’Europe occidentale. 1. Calanoïdes et

Harpacticoïdes. Boubée, 1967. ([115])

● B. Dussart, Les Copépodes des eaux continentales d’Europe occidentale...: Cyclopoïdes et biologie, vol.

2. N. Boubée et Cie, 1969. ([116])

● P. D. N. Hebert, “The Daphnia of North America: an illustrated fauna,” CD-ROM, Univ. Guelph, 1995.

([117])

● N. N. Smirnov, Cladocera: the Chydorinae and Sayciinae (Chydoridae) of the world, vol. 11. SPB

Academic Pub., 1996. ([118])

● J. W. Reid and H. Ueda, Copepoda: Cyclopoida: Genera Mesocyclops and Thermocyclops. Backhuys

Publishers, 2003. ([119])

● A. Petrusek, F. Bastiansen, and K. Schwenk, “European Daphnia species (EDS)-taxonomic and genetic

keys.[Build 2006-01-12 beta],” CD-ROM Distrib. by authors. Dep. Ecol. Evol. JW Goethe-University,

Frankfurt am Main, Ger. Dep. Ecol. Charles Univ. Prague, Czech Repub., 2005. ([120])



Appendix II-E

I. E. McCauley, “The estimation of the abundance and biomass of mesozooplankton in samples,” in A

manual on methods for the assessment of secondary productivity in fresh waters, 1984, pp. 228–265

[49], including the following meta-analysis:

a. H. J. Dumont, I. Van de Velde, and S. Dumont, “The dry weight estimate of biomass in a

selection of Cladocera, Copepoda and Rotifera from the plankton, periphyton and benthos

of continental waters,” Oecologia, vol. 19, no. 1, pp. 75–97, 1975. ([67])

b. H. Bottrell et al., “A review of some problems in mesozooplankton production studies,” Nor.

J. Zool., vol. 24, pp. 419–456, 1976. ([48])

c. M. L. Pace and J. D. Orcutt, “The relative importance of protozoans, rotifers, and crustaceans

in a freshwater mesozooplankton community,” Limnol. Oceanogr., vol. 26, no. 5, pp. 822–

830, 1981. ([70])

d. R. A. Rosen, “Length-dry weight relationships of some freshwater mesozooplanktona,” J.

Freshw. Ecol., vol. 1, no. 2, pp. 225–229, 1981. ([71])

e. G. Persson and G. Ekbohm, “Estimation of dry-weight in mesozooplankton populations -

Methods applied to Crustacean populations from lakes in the Kuokkel area, Northern

Sweden,” Arch. fur Hydrobiol., vol. 89, no. 1–2, pp. 225–246, 1980. ([68])

f. C. W. Burns, “Relation between filtering rate, temperature, and body size in four species of

Daphnia,” Limnol. Oceanogr., vol. 14, no. 5, pp. 693–700, 1969. ([69])

g. M. J. Burgis, “Revised estimates for the biomass and production of mesozooplankton in Lake

George, Uganda,” Freshw. Biol., vol. 4, no. 6, pp. 535–541, 1974. ([121])

h. T. R. Jacobsen and G. W. Comita, “Ammonia-nitrogen excretion in Daphnia pulex,”

Hydrobiologia, vol. 51, no. 3, pp. 195–200, 1976. ([122])

i. E. Michaloudi, “Dry weights of the mesozooplankton of Lake Mikri Prespa (Macedonia,

Greece),” Belgian J. Zool., vol. 135, no. 2, pp. 223–227, 2005. ([97])

II. L. A. Smock, “Relationships between body size and biomass of aquatic insects,” Freshw. Biol., vol. 10,

no. 4, pp. 375–383, 1980. ([123])

III. D. A. Culver, M. M. Boucherle, D. J. Bean, and J. W. Fletcher, “Biomass of Freshwater Crustacean

Zooplankton from Length–Weight Regressions,” Can. J. Fish. Aquat. Sci., vol. 42, no. 8, pp. 1380–

1390, Aug. 1985. ([124])

IV. J. Watkins, L. Rudstam, and K. Holeck, “Length-weight regressions for mesozooplankton biomass

calculations – A review and a suggestion for standard equations.,” Cornell Biol. F. Station. Dep. Nat.

Resour., 2011. ([125])



Appendix III: Microbial Plankton

N.A.

Appendix IV: Periphyton

N.A.

Appendix V: Water Chemistry

N.A.



Appendix VI: High Frequency Measurements

Appendix VI-A: General Procedures for Calibration of Field Thermometers

To calibrate field thermometers when a commercial refrigerated water bath is available (Paraphrased from

[6]):

1. Precool the sensor of the thermometer/thermistor being checked (field thermometer) to 0°C by immersing

it in a separate ice/water bath.

2. Immerse the field and calibration temperature sensors in the refrigerated bath with a water temperature

of approximately 0°C.

3. Position the temperature sensor(s) so that they are properly immersed and the scales can be read. Stir the

water bath and allow at least 2 minutes for the thermometer readings to stabilize.

4. Without removing the temperature sensor from the refrigerated water bath, read the field thermometer(s)

to the nearest graduation (0.1 or 0.5°C) and the calibration thermometer to the nearest 0.1°C.

a. Take three readings within a 5-minute span for each field thermometer.

b. Calculate the mean of the three temperature readings for each field thermometer and compare

its mean value with the calibration thermometer.

c. If a liquid-filled field thermometer is found to be within ±1 percent of full scale or ±0.5°C of the

calibration thermometer, whichever is less, set it aside for calibration checks at higher temperatures.

d. If a field thermistor is found to be within ±0.2°C of the calibration thermometer, set it aside for

calibration checks at higher temperatures. Troubleshooting steps must be taken. If troubleshooting

fails, the sensor should be returned to the manufacturer for proper calibration, repair, or

replacement [5].

5. Repeat steps 1–4 in 25°C and 40°C water. Keep the bath temperature constant. Check the thermistors at

two or more additional intermediate temperatures (for example, 15°C and 30°C).

6. Tag the acceptable and calibrated thermometers/thermistors with calibration date and certifier’s initials.

To calibrate field thermometers when a commercial refrigerated water bath is not available (Paraphrased

from [6]):

For the 0°C calibration

1. Freeze several ice cube trays filled with deionized water.

2. Fill a 1,000-milliliter (mL) plastic beaker or Dewar flask three fourths (3/4) full of crushed, deionized ice.

Add chilled, deionized water to the beaker. Place the beaker of ice/water mixture in a larger, insulated

container or Dewar flask. Place the calibration thermometer into the ice/water mixture and make sure that



the temperature is uniform at 0°C by stirring and checking at several locations within the bath.

3. Precool the sensor of the field thermometer(s) to 0°C by immersing in a separate ice/water bath.

4. Insert the field thermometer(s) into the ice/water mixture. Position the calibration and field thermometers

so that they are properly immersed, and the scales can be read. Periodically stir the ice/water mixture and

allow at least 2 minutes for the thermometer readings to stabilize.

5. After the readings stabilize, compare the temperature of one field thermometer at a time with that of the

calibration thermometer. Without removing the temperature sensor(s) from the test bath, read the field

thermometer(s) to the nearest graduation (0.1 or 0.5°C) and the calibration thermometer to the nearest

0.1°C.

a. Take three readings for each thermometer within a 5-minute span.

b. Calculate the mean of the three temperature readings for each thermometer and compare its

mean value with the calibration thermometer.

c. If the field liquid-filled thermometer is found to be within ±1 percent of full scale or ±0.5°C of the

calibration thermometer, whichever is less, set it aside for calibration checks at higher temperatures.

d. If the field thermistor is found to be within ±0.2°C of the calibration thermometer, set it aside for

calibration checks at higher temperatures.

For the “room temperature” calibration (25°C )

1. Place a Dewar flask or container filled with about 3.8 liters (1 gallon) of water in a box filled with packing

insulation. (A partially filled insulated ice chest can be used for multiparameter instruments.) Place the

calibration container in an area of the room where the temperature is fairly constant (away from drafts,

vents, windows, and harsh lights).

2. Properly immerse the calibration and field thermometer(s) in the water. Cover the container and allow the

water bath and thermometers to equilibrate.

3. Stir the water and, using the calibration thermometer, check the bath for temperature uniformity. Repeat

this every 2 hours. It may be necessary to let the bath equilibrate overnight.

4. Compare one field thermometer at a time against the calibration thermometer, following the procedures

described above in step 5 for the 0°C calibration.

For each temperature that is greater than 25°C

1. Warm a beaker of 1,000 mL or more of water to the desired temperature (for example, 40°C) and place it

on a magnetic stirrer plate.

2. Follow the procedures described above in step A5 for the 0°C calibration.



Appendix VI-B: General Procedures for Calibration of pH Sensors

Summary of calibration process for pH meters, including a wide range of available equipment, is compiled

from Wagner and colleagues (2006) [5] and Ritz and Collins (2008) [8] as follows:

1. Rinse the pH sensor, thermistor or thermometer, and calibration cup with pH-7 buffer solution.

2. Pour the fresh pH-7 buffer solution into the rinsed calibration cup. Immerse the sensor/probe into

the solution, making sure the sensor’s glass bulb is in solution by at least 1 cm.

● Allow the instruments to equilibrate for at least 1 minute for temperature equilibration

before proceeding.

3. Measure and record the temperature, pH, and associated millivolt reading (if available), along with

lot numbers and expiration dates of the pH buffers. This standardization process is repeated with

fresh pH-7 buffer solution until two successive values of the temperature-adjusted pH-7 readings are

obtained.

4. Rinse the pH sensor, thermistor or thermometer, and calibration cup with de-ionized water. Repeat

the standardization process with a pH-4 or pH-10 buffer solution to establish the response slope of

the pH sensor.

5. Use for instance, pH-4 or pH-10 buffer to establish the slope of the calibration line at the temperature

of the solution.

✓ A buffer that brackets the expected range of pH values in the environment should be

selected.

6. Record the second temperature-corrected pH value, temperature, millivolt readings, lot numbers,

and expiration dates. Rinse the pH sensor, thermistor or thermometer, and calibration cup are with

de-ionized water again.

7. The pH-7 buffer solution is then used to rinse, fill, and check the pH-7 calibration measurement. If

the pH sensor reading is 7 ±0.1 pH units (as a correctly calibrated pH sensor can accurately measure

pH to ±0.2 pH unit, See Table 8-1) the slope adjustment has not affected the calibration. If the

accuracy standard is not met, the calibration and slope adjustment steps must be repeated.

✓ If re-calibration and troubleshooting steps fail, the pH sensor or monitoring probe must be

replaced.

8. Once the slope-adjustment step is completed satisfactorily, the third buffer solution can be used as

a check for calibration range and linearity of electrode response.

● The temperature and pH values are read and recorded along with the lot numbers and

expiration dates of the pH buffers; however, the ±0.1 pH accuracy should not be expected to

be achieved over the full range from pH-4 to pH-10 for a monitoring sensor. The third buffer

should be within ±0.2 pH unit value [5].



Appendix VI-C: General Procedures for Calibration of DO Sensors

Three steps to calibration, as compiled from Wagner and colleagues (2006) [5] and Rounds and colleagues

(2013) [9], are as follows:

Step 1. Calibration of a DO meter at 100-percent oxygen saturation is made by adjusting the meter reading

for air saturated with water vapor to a value obtained from a DO solubility table. One or two-points

calibration can be carried out to ensure 100% saturation oxygen environment, followed by calibration at zero

DO.

✓ The two-points calibration adds complexity to the calibration process and is not recommended by all

manufacturers of optical sensors. Be sure first to understand the instrument capabilities and then

determine the best course of action for your field work.

Step 2. A zero-DO sensor-performance check should be carried out. For that reason, the zero-DO sodium

sulfite solution4 should be prepared.

✓ Before immersing sensor in the zero-DO solution, the wiper (if applicable) should be removed

from the unit to avoid saturating it with the sodium sulfite solution. Then, the sensor and the

wiper should be rinsed thoroughly and then reinstalled.

✓ Multiple and thorough rinses with deionized water are necessary to restore the sensor to

proper operating condition and prevent bias to subsequent measurements.

Step 3. Calibration of DO sensors: Air-calibration chamber in air, calibration with air-saturated water, air-

calibration with a wet towel and air calibration chamber in water methodologies were explained in details in

[9]. As mostly recommended by the manufacturers for optical sensor calibration, calibration with air-

saturated water method is explained in details in this SOP.

Calibration with air-saturated water:

This method is considered to provide the best accuracy for calibration of optical sensors. The calibration

should be done while the instrument is kept in water that is saturated with oxygen at a known

temperature and ambient atmospheric pressure.

1. Required Equipment (as adapted from [9]):

a. 5-gallon (around 19 liters) bucket, three-quarters full of tap water, or manufacturer-provided

aeration chamber

b. 10-gallon (around 38 liters) aquarium air pump with two outlets

c. 10-foot-length (around 300 cm) of aquarium pump tubing

4 The zero-DO sodium sulfite solution: Dissolve 1 gram sodium sulfite in 1 liter of deionized water (0.008 M solution, prepared fresh just before the field trip or during week of use).



d. 2 X Gas-diffusion (air) stones

2. The methodology (as adapted from [9]):

1. Attach the pump tubing to the pump and then the two air stones to the ends of the tubing.

Place the tubing with air stones at the bottom of the filled bucket.

2. Turn on the pump and aerate the water for a minimum of 30 minutes.

✓ The pump could be left to operate continuously (24/7) in order to have a ready supply

of air-saturated water on hand for calibration in the laboratory or for transport and

calibration in the field.

3. Take care to keep air bubbles off the optical sensor (the luminescence tip or membrane).

4. Place the DO sensor (or multi-parameter sensor) in the bucket and allow 5 to 10 minutes for

the sensor to come to thermal equilibrium with the aerated water. Take care not to place

the sensor over or in the bubbles from the air stone.

5. Read and record the temperature of the calibration water to the nearest 0.1 °C.

6. Using a calibration-checked altimeter-barometer, determine the ambient atmospheric

pressure to the nearest 1 mm of mercury.

7. Using oxygen-solubility table, as described in [9]. Determine the DO saturation value at the

measured temperature and atmospheric pressure of the calibration water.

✓ An interactive program is also available for producing a table of DO saturation

values to the nearest 0.1 or 0.01 mg/L over user-defined ranges of temperature

and barometric pressure and a table of salinity correction factors over user-

defined ranges of specific conductance

(http://water.usgs.gov/software/dotables.html).

8. Salinity Corrections:

✓ A reliable pocket altimeter can be used to measure uncorrected (true)

barometric pressure to the nearest 1 millimeter (mm) of mercury; a specific

conductance meter can be used to measure salinity.

9. Calibration of field barometers should be checked before each field trip, preferably by

checking with an official weather station [9].

10. Verify that the instrument reading is within ±0.2 mg/L or 2 percent of the computed

saturation value. Alternatively, use more stringent accuracy criteria that reflect the data-

quality requirements of the study. If the field calibration or calibration check fails to meet

the established criterion,

a. use a different instrument (if available), and

b. Do not collect or report data using an instrument that has failed calibration.

11. Record calibration information in instrument log books and transfer calibration data into

electronic records or onto paper field forms at the time of calibration.

http://water.usgs.gov/software/dotables.html)



Appendix VI-D: General Procedures for Calibration of Conductivity Sensors

The general procedures, that apply to most of the instruments used in conductivity measurements, described

in [10] and [5] are summarized below:

1. Calibration is performed at the field site with calibration standard solutions that have been allowed

to equilibrate to the temperature of the water being monitored.

2. The instrument is turned on. Allow sufficient time for electronic stabilization.

3. Select the correct amount of conductivity standard solutions that will bracket the expected sample

conductivity. Equilibrate the standards and the conductivity sensor to the temperature of the sample.

✓ Verify that the date on the standards has not expired.

✓ Put bottles of standards in a minnow bucket, cooler, or large water bath that is being filled with

ambient water.

✓ Allow 15 to 30 minutes for thermal equilibration. Do not allow water to dilute the standard.

4. Accurate conductivity measurements depend on accurate temperature measurements or accurate

temperature compensation. Rinse the conductivity sensor, the temperature meter (calibrated

thermistor or a hand-held device), and a container large enough to hold the dip-type sensor and the

thermometer.

● First, rinse the sensor, the thermometer, and the container three times with deionized water.

● Next, rinse the sensor, the thermometer, and the container three times with the standard to be

used.

5. Immerse the sensor/probe (and the thermometer) into the rinsed calibration cup and pour in fresh

calibration standard (solution). Allow at least one minute for temperature equilibration before

proceeding.

6. Measure water temperature.

7. If applicable, initiate the Calibration Software and follow the instructions of the manufacturer to

complete calibration. If not, continue with Step 9.

8. Whisk the conductivity sensor up and down under the solution surface to expel air trapped in the

sensor. Read the instrument display. Agitate the sensor up and down under the solution surface

again and read the display. Repeat the procedure until consecutive readings are the same.

9. Record the instrument reading and adjust the instrument to the known standard value.

✓ If an instrument cannot be adjusted to a known calibration standard value, develop a calibration

curve. After temperature compensation, if the percentage difference from the standard exceeds

5 percent, refer to the troubleshooting section.

10. Record following parameters in the instrument log book and on field forms:

● The temperature of the standard solution.

● The known and the measured conductivity of the standard solution (including ± variation).

● The temperature-correction factor (if necessary).



11. Discard the used standard into a waste container. Thoroughly rinse the sensor, thermometer, and

container with deionized water.

12. Repeat steps 7 through 11 with the second conductivity standard.

✓ The purpose for measuring a second standard is to check instrument calibration over the range

of the two standards.

✓ The difference from the standard value should not exceed 5%.

✓ If the difference is greater than 5%, repeat the entire calibration procedure. If the second reading

still does not come within 5%of standard value, refer to the troubleshooting section or calibrate

a backup instrument.

Appendix VI-E: General Procedures for Calibration of in situ Fluorescent Sensors

The general procedures, that apply to most of the instruments used in situ chlorophyll measurements are

summarized below:

1. Pour the correct amount of clear deionized or distilled water into the calibration cup.

2. Immerse the probe in the water.

3. Let the probe to get stabilized (around 40 seconds to 1 minute).

4. Observe and record the reading. Remove the probe out of water.

5. Pour the required amount of the standard solution with the known content/ratio into the calibration

cup.

6. Place the sensor into the solution.

7. Let the probe to get stabilized (around 40 seconds to 1 minute).

8. Observe and record the reading. Remove the probe out of the standard solution.

9. Rinse the probe/sensor in tap/purified water. Leave the sensor to dry.

Appendix VI-F: General procedures for 3-points calibration of ammonium and nitrate sensors

● The commonly followed calibration procedure for nitrate is identical to the procedure for

ammonium, except that the calibration standard solution values are in mg/L NO3 - -N instead

of NH4+ -N.

1. Pour a sufficient amount of 1 mg/L NH4 + -N (or 1 mg/L NO3 - -N) calibration standard solution at

ambient temperature in a clean and dry or pre-rinsed calibration cup.

2. Carefully immerse the sensor into the solution, making sure the sensor’s tip is in solution by at least

1 cm.

3. Allow at least 1 minute for temperature equilibration before proceeding.

4. Observe and record the readings when they are stable (i.e. data shows no significant change for

approximately 40 seconds).



5. Rinse the sensors in deionized water between changes of the calibration solutions.

6. Pour a sufficient amount of 100 mg/L of NH4 + -N (or 100 mg/L NO3 - -N) calibration standard

solution at ambient temperature into a clean, dry or pre-rinsed calibration cup.

7. Carefully immerse the sensor into the solution.

8. Allow at least another 1 minute for temperature equilibration before proceeding.

9. Observe and record the readings when they are stable (i.e. data shows no significant change for


10. Rinse the sensors in deionized water between changes of the calibration solutions.

11. Immerse the sensor in the pre-chilled 1 mg/L NH4 + -N (or 1 mg/L NO3 - -N) calibration standard

solution ensuring that the temperature is at least 10°C different than ambient.

12. Allow at least 1 minute for temperature equilibration before proceeding.

13. Observe and record the readings when they are Stable (i.e. data shows no significant change for


14. Rinse the sensor in tap or purified water.

Appendix VII QA &QC

N.A.

deliverable no 4.1: standard operating protocol (sop) on

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