spi

Available online at www.sciencedirect.com

Bioresource Technology 99 (2008) 3949–3964

Review

Microalgae immobilization: Current techniques and uses

Ignacio Moreno-Garrido *

Institute of Marine Sciences of Andalucıa (CSIC), Campus Rıo San Pedro, s/n 11510, Puerto Real, Cadiz, Spain

Received 19 July 2006; received in revised form 23 May 2007; accepted 23 May 2007Available online 9 July 2007

Abstract

Information about advances in immobilization techniques and biotechnology use of freshwater and marine microalgae is scattered.This work aims to bring together the main recent research about the topic. Passive and active immobilization techniques used on mic-roalgae are listed and described in the text. Effect of immobilization on growth and metabolism of the cells is also reviewed. Current usesof immobilized microalgae include metabolite production, culture collection handling, obtaining of energy and removing of undesired orvaluable substances from media (nutrients, metals and different pollutant agents). Applications of immobilized microalgae in environ-mental aquatic research have been recently increased: novel immobilization techniques as well as the use of living microalgae as biosen-sors in electronic devices designed to measure toxicity of substances and effluents demonstrated to be a very promising topic inbiotechnology research. Recent research pointed out the advantages of mixed bacterial–algal co-immobilized systems in water treatmentplants. Application of immobilized systems to the production of non-contaminant energy (as H2 obtained from algal cultures) is also animportant topic to be explored in the next years.� 2007 Elsevier Ltd. All rights reserved.

Keywords: Microalgae; Phytoplankton; Immobilization; Biotechnology

1. Introduction

Microalgae (sensu lato, it means including prokaryoticphotosynthetic microorganisms such as cyanobacteria)are organisms that play a key role in aquatic ecosystems.It is estimated that around 40% of global photosynthesisis performed by microalgae (Falkowsky, 1980). Microalgaeform the basis of most of trophic aquatic chain. In somecoastal environments, biomass of microphytobenthos canmatch or even exceed the biomass of bacteria (La Rosaet al., 2001). The use of microalgae in biotechnology hasbeen increased in recent years, these organisms being impli-cated in food, cosmetic, aquaculture and pharmaceuticalindustries (Borowitzka and Borowitzka, 1988), but smallsize of single cells implies a problem in the application ofbiotechnology processes to those organisms. Cell immobi-lization techniques have been developed in order to solve

0960-8524/$ - see front matter � 2007 Elsevier Ltd. All rights reserved.

doi:10.1016/j.biortech.2007.05.040

* Tel.: +34 956832612; fax: +34 956834701.E-mail address: [email protected]

those problems. The use of immobilized algal cells in waterpurification processes has been reported from long ago(Robinson et al., 1988), as microalgae form part of theorganisms fixed in percolating filters of wastewater treat-ment plants. But at the end of the sixties of the past cen-tury, novel techniques for immobilizing biocatalysts ingeneral (from enzymes to whole cells) began to spread inthe literature (Papageorgiou, 1987), and the use of immobi-lization techniques diversifies. Immobilized algae have beenused for biomass obtain and macronutrient removal. Theextremely high accumulation capacity of some of theseorganisms for potentially dangerous substances (Maedaand Sakaguchi, 1990) has been also exploited for bioreme-diation techniques applied on polluted waters (speciallyinvolving metals: Greene and Bedell, 1990). This capacityhas also been exploited in order to pre-concentrate thesesubstances and thus facilitate the measurement of tracesin environment (Carrilho et al., 2003; Singh and Prasad,2000). Lower growth rates of immobilized algal cells, prob-ably due to restrictions in the diffusivity of nutrients to theimmobilized cells, as well as protection of cells entrapped in

mailto:[email protected]

3950 I. Moreno-Garrido / Bioresource Technology 99 (2008) 3949–3964

immobilizing matrixes can also be exploited in order tofacilitate handling of culture collections (Lukavsky,1988). Environmental uses of immobilized algal cells arenot restricted to pollutant removal. These techniques havebeen recently used in field toxicity measurement experi-ments (Admiraal et al., 1999; Moreira dos Santos et al.,2002, 2004; Moreira et al., 2006). An important part ofthe current work in the field of the toxicity measurementis, however, focused in the design of microalgal-based bio-sensors (Chouteau et al., 2004; Podola et al., 2004; Shit-anda et al., 2005). Recent uses of microalgal technologyinvolving production of hydrogen (Dante, 2005; Kapdanand Kargi, 2006) or electricity (Kadam, 2002) could beimproved by the use of immobilization techniques. Verywell known reviews on immobilization techniques and usesfor algae date from the end of the eighties (Robinson et al.,1986; Codd, 1987; Papageorgiou, 1987). Cassidy et al.(1996) made a revision of the environmental applicationof immobilized cells (not only algae). Jen et al. (1996) sum-marized in a review the new techniques till that year onhydrogels for cell immobilization in general. One of themost recent reviews on the topic is the work of Mallick(2002), centred in the use of immobilized algae for waste-water, nitrogen, phosphorus and metal removal. But abig amount of information about all these and other usesfor immobilized cells still remains scattered. The purposeof this review is, thus, to bring together all recent informa-tion to date about microalgal immobilization techniquesand the current uses for this biological material, in orderto facilitate researchers the task of finding referencesrelated to their work in this field.

2. Immobilization techniques for microalgal cells

Most of the immobilization techniques devised formicroorganisms in general can be applied to microalgae,with the limitation of light transmission if living cells areintended to be immobilized. Immobilization techniquescan be primarily divided into two groups: ‘‘passive’’ and‘‘active’’ immobilization.

2.1. Passive immobilization

Many microorganisms (including some groups of micro-algae) have a natural tendency to attach to surfaces andgrow on them (Robinson et al., 1986). This characteristiccan be exploited in order to immobilize cells on carriersof different types (Codd, 1987). Normally, those processesare easily reversible and contamination of effluents withunstuck cells is unavoidable. Adsorbent materials (carriers)for passive immobilization can be natural or synthetic.With respect to natural carriers, recently efforts have beenmade involving loofa biomass. Loofa sponges are thefibrous support of the fruit of different species from thegenus Luffa (L. cylindrica – possible synonym: L. aegypti-aca, L. operculata, L. acutangula). The sponge is obtainedfrom dry fruits after removing the pericarp tissue. This car-

rier is non-toxic and (it is said to be) non-reactive, cheap,mechanically strong and stable in long-term cultures (Liuet al., 1998). Akhtar et al. (2004) used loofa sponge bio-mass in order to immobilize cells of Chlorella sorokiniana,to remove nickel(II) from aqueous solutions. This immobi-lized system demonstrated to accumulate 25% more nickelthan free cells after 20 min exposition. This immobilizingtechnique using loofa sponges has also been used for fun-gus such as Phanaerochaete chrysosporium (Iqbal andEdyvean, 2004, 2005; Ahmadi et al., 2006) in wastewatertreatments; bacteria as Zymomonas nobilis (Vignoli et al.,2006) for sorbitol production; and microbial bio-systems(Nagase et al., 2006) for degradation of carbendazim and2,4-dichlorophenoxyacetic acid. For cells that have no nat-ural trend to attach this type of support, Ogbonn et al.(1996) reported the possibility of the co-use of chitosanin order to increase the flocculating process of free cellsover the loofa surfaces. Liu et al. (1998) compared the cel-lular adsorption capacity of loofa sponge cubes and poly-urethane cubes, and found no differences for plant(Coffea arabica) free cells. A problem on designing researchinvolving loofa sponge biomass is repeatability. Structureof the skeleton of fruits varies from a plant to another infunction of culturing conditions: each loofa sponge has dif-ferent structure (Liu et al., 1998). In any case, for industryor commercial purposes, passive immobilization of algalcells on loofa sponges seems to be a very promising field.

Synthetic materials are also widely used in experimentsinvolving passive immobilization. Urrutia et al. (1995)immobilized Scenedesmus obliquus cells in polyvinyl andpolyurethane in order to remove nitrate from water. Sur-vival of adsorbed cells was compared with entrapped cells(it means, cells immobilized by ‘‘active’’ immobilization),by mixing concentrated cells with one of the pre-polymers.Cellular growth is higher for adsorbed cells than that mea-sured for entrapped cells, possibly due to the toxicity of thepre-polymers (although these authors reported that notoxic effects were found due to the residual presence ofpre-polymer reagents). Yamaguchi et al. (1999) achieveda noticeable degradation of hydrocarbons by the colorlesshydrophobic microalgae Prototheca zopfii, adsorbed to8 mm-cubes of polyurethane foam in a bubble reactor.Archambault et al. (1990) described a reactor where plantcells attach by natural adhesion to short-fiber polyestermaterial named 7607.

Attachment of periphyton to different surfaces can alsobe used in ecology studies. Admiraal et al. (1999) usedmicrobenthic algae and bacteria colonizing glass discs tomeasure the response of these organisms to different levelsof metal pollution. Danilov and Ekelund (2001) comparesettlement patterns of periphyton on glass, wood and plas-tic in lakes of different trophic status, this order (glass >wood > plastic) being the preference of the periphytic spe-cies to attach on. Studies on glass attached periphyton havealso been used to measure the influence of water current instreams on the structure of colonizing periphytic algae(Ghosh and Gaur, 1998). Periphyton attachment to glass

I. Moreno-Garrido / Bioresource Technology 99 (2008) 3949–3964 3951

slides (mainly diatoms) and population growth on this typeof surfaces has been studied by Brandini et al. (2001) in asubtropical estuary during a whole year, permittingresearchers to compare differences caused by depth, light,temperature and grazing pressure. Nayar et al. (2005) alsostudied the settlement of periphytic algae in glasses in atropical estuary (Singapore), characterizing production interms of 14C uptake. Following those authors, planktonicdiatom species such as Skeletonema costatum and Thalass-

iosira rotula, together with cyanophytes such as Synecho-

coccus sp., dominated the assemblages. Glass-immobilizedfilaments of Anabaena sp., have also been used in hydrogenproduction experiments (Robinson et al., 1986). These sup-port materials can be only used with cells from speciesshowing natural trend to attach o aggregate (mainly dia-toms and cyanophytes), if the material is not previouslytreated or a co-flocculant is not used. Other carriers suchas Biolite� (a ceramic material) have been used for immo-bilizing bacteria (Prieto et al., 2002), but no referencesrelated to photosynthetic microorganisms have been found.

2.2. Active immobilization

Regarding active immobilization techniques, the use offlocculant agents, chemical attachment and gel entrapmentshould be distinguished.

2.2.1. Flocculant agents

Flocculant agents were primarily used in order to avoidtedious and expensive centrifugation when algae areintended to be removed from a liquid medium. Amongthe commonly used flocculants, chitosan has been the mostwidely employed. Chitosan is a linear amino polysaccha-ride of b-D-glucosamine (2-amino-2-deoxy-b-D-glucan)units joined by (1! 4)-linkages (Oungbho and Muller,1997). It is obtained through a chitin (obtained from exo-skeletons of crustaceans) alkaline deacetylation. This poly-saccharide presents positive-charged amino groups,providing very interesting properties for adsorbing nega-tive-charged particles and it is demonstrated to be usefulfor a large number of microalgal species (Lubian, 1989).This substance is biodegradable, and thus can be used inharvesting of algae for nutritional purposes. Largeamounts of Euglena gracilis have been removed frommedia by Gualtieri et al. (1988), reaching a 96–98% reduc-tion of suspended cells with 200 mg L�1 of chitosan at pH7.5. The inconvenience of chitosan in immobilizing tech-niques is its weak stability. Kaya and Picard (1996) triedto solve this problem using high viscosity chitosan andkonjac flour (glucomannan, obtained from tuberous rootof the konjac tree – Amorphophalus konjac) in order toenhance the stability of floccules immobilizing viable cellsof Scenedesmus bicelularis to use them in tertiary treatmentof wastewaters, but concluded that konjac flour did not sig-nificantly modify the rheological properties of mixed chito-san solutions. High viscosity chitosan gels (2% p/v) showeda higher chemical stability in the experiments described by

these authors. It seems that the content of phosphateanions in the media helps to maintain the stability of chito-san gels. Chitosan can interfere in the growth of immobi-lized algae: Moreira et al. (2006) found low growth rates(increasing factor not higher than 4 after 3 days) of Phae-

odactylum tricornutum immobilized in alginate treated withchitosan as additional hardener, while cells immobilized inalginate beads hardened only with CaCl2 or SrCl2 showedincreasing factors of 31 and 76 times, respectively.

2.2.2. Chemical attachment

Chemical attachment presents some great disadvantageswhen living cells are intended to be immobilized, becausethe chemical interaction (mainly due to covalent bonding,cross-linking – involving glutaraldehyde, for instance – orphotocrosslinkable resins) causes damages in cellular sur-face and drastically reduces viability of cells. Ion attractionis not so harmful to living organisms, but effectiveness ofthis technique depends on pH and ionic strength of the sur-rounding media (Codd, 1987). It is necessary to remindthat pH in the immediate medium of a living microalgaecan reach very high values in light, due to photosyntheticmetabolism. Nevertheless, some experiments do not requireactive metabolism of cells, and non-living organisms areused in chemical attachment immobilization techniques.Thus, Seki and Suzuki (2002) compared the adsorptioncapacity of two floc-type biosorbents in order to removean ‘‘inactivated’’ marine microalga which is able to accu-mulate Cd and Pb from aqueous media. The species used,Heterosigma akashiwo, forms red tides in coasts of Japan,thus being an inexhaustible and attractive material forthe biosorbent, following those authors. The two materialscompared were milk casein floccules (used as an immobiliz-ing flocculant protein and as a biosorbent at the same time)and glutaraldehyde. The technique developed comprised asmuch as 67% of microorganisms present in the cultures.

2.2.3. Gel entrapment

This method is the most widely used technique for algalimmobilization. Following Codd (1987), gel entrapmentcan be performed by the use of synthetic polymers (acryl-amide, photocrosslinkable resins, polyurethanes), proteins(gelatine, collagen or egg white) or natural polysaccharides(agars, carrageenans or alginates).

2.2.3.1. Synthetic polymers for gel entrapment. Blanco et al.(1999) described the use of polysulphone (a thermoplasticmaterial) and epoxy resin entrapping cells of the cyanobac-teria Phormidium laminosum, in order to check the capacityof biosorption (and desorption, by the use of 0.1 M HCl) ofCu(II), Fe(II), Ni(II) and Zn(II). Epoxy resins consist oftwo components that react with each other forming a hard,inert material. One of them is a bisphenol and the other isepichlorohydrin. Toxicity of the components makes epoxyresins not suitable for entrapping living cells. In the exper-iments described by Blanco et al. (1999), cells were previ-ously dried (and thus killed) at 55 �C. Authors concluded


that metal accumulation in the beads directly depends onthe entrapped cyanobacterial biomass. The same cyano-bacterial species has been used by Garbisu et al. (1991) inexperiments designed in order to remove nitrate from waterby cells immobilized in polyvinyl and polyurethane foams.In the described case, polyvinyl was hydrophilic (PV-50)and the polyurethane was prepared from Hypol FHP2002, a special grade of polyether polyisocyanate pre-poly-mer that is mixed with a hydroxyl group. Entrapment rap-idly leads to the death of cells, and only slow passivecolonization of foams was used for experiments involvingnitrate removal. These authors also found that recentlymade foams were toxic to cells and inhibited even passivecolonization, possibly due to the leftover pre-polymers.Toxicity disappears when foams were pre-autoclaved. Por-

phyridium cruentum cells have been immobilized in polyure-thane foam by Thepenier et al. (1985) in order to producepolysaccharides. After immobilization, these authors didnot detect any oxygen evolution, due to the high degreeof cellular destruction during the immobilization reaction.At least four days had to pass before the immobilized col-onies began to grow (after intensive rinses). The genus Por-phyridium produces high amounts of exo-polysaccharides,which could protect inner cells in colonies or cellulargroups from toxicity of pre-polymers. The same couldoccur with mucilaginous cyanobacteria (Streble and Kra-uter, 1987), protected by sheaths. These sheaths can stickone to another forming colonies of filaments. Thus, it ispossible that inner filaments in a colony could survive totoxicity of the pre-polymers of polyurethane foams andafter rinsing of pre-polymer leftovers, and then begin togrow and colonize the foams. In the experiment of Thepe-nier et al. (1985), after 10 days, oxygen production in batchcultures began to decrease dramatically and stabilized atlow values on day 30, but polysaccharide production beganto increase from day 40. This foam also permitted therelease of cells to the media. Kawabata et al. (1990) suc-cessfully immobilized bacterial cells (Escherichia coli) onthe surface of cross-linked poly(N-benzyl-4-vinylpyridi-nium bromide) containing styrene (BVPS). The cellsremained alive and were able to produce L-aspartic acidfrom ammonium fumarate. Although this technique hasnot been applied to microalgae, it seems to be suitablefor immobilizing living algal cells. Willke et al. (1994) alsosuccessfully immobilized the bacteria Paracoccus denitrifi-

cans in polycarbamoylsulphonate (PCS), a very low toxichydrogel matrix. Survival of cells to immobilization proce-dure was greater than 99%. Rarely, references concerningimmobilization with this technique can be found, but itcould be a very suitable technique in order to avoid the sta-bility problems of alginate gels (over all in marine environ-ments, as will be further discussed). Other techniques, asthose described in Jeon et al. (2002), imply polyvinyl alco-hols and glutaraldehyde. This technique does not seem tobe recommendable if living cells are intended to be immo-bilized. Silica gels can be used for immobilizing microalgalcells. Rangasayatorn et al. (2004) described a bioassay

involving Spirulina platensis in order to check cadmiumadsorption of immobilized cells in alginate and silica gel.The latter technique is also called sol–gel technique, andit is described in Weller (2000) for immobilizing antibodies.Gelation of the sol occurs when pH of alkali silicate is car-ried under values of 10. But in the case described byRangasayatorn et al. (2004), particles of silica gel obtainedafter crushing were dried at 80 �C, and thus survival of cellsis not a target point of the experiment. Nevertheless, cad-mium adsorption of silica gel entrapping cells is as highas in the alginate entrapped cells (near 100% of Cd adsorp-tion after 1 h exposition, 10 mg L�1 being the initial Cdconcentration). Overnight drying (80 �C) of polysilicatematrix seems to be a common procedure, as it is also usedby Stark and Rayson (2000) in a work comparing metal–ion binding capacity of different immobilized materials(Sphagnum peat, top soil, peat, compost peat, organic peat,peat replacer, dead Chlorella vulgaris cells and cells fromDatura innoxia – a solanacean plant). In this case, biomassfrom Chlorella was obtained by a commercial algae-basedbiosorbent (Algasorb�), and this material shown to be veryefficient in metal removal (but surpassed by peat andorganic compost peat) (Singh and Prasad, 2000).

2.2.3.2. Proteins for gel entrapment. Proteins are not widelyused in microalgal immobilization. In the reviews of Rob-inson et al. (1986) and Papageorgiou (1987), immobiliza-tion by proteins does not appear among the techniquesused. Only Codd (1987) mentioned egg white, collagenand gelatine as examples of possible immobilization tech-niques involving proteins. Other organisms apart frommicroalgae have been immobilized by these techniques(yeasts in hen egg white, for instance: Kubal and D’Souza,2004). The use of glutaraldehyde as a cross-linker for eggwhite does not predict good results with living microalgae,but could be used for dead cells.

2.2.3.3. Natural polysaccharides for gel entrapment. Gelentrapment in natural polysaccharide matrixes is the mostwidely used immobilization technique for microorganismsin general (and microalgae in particular). Among them,carrageenan, agar and alginate are the most employed.

Carrageenan is a collective term for polysaccharides pre-pared by water alkaline extraction from some Rhodophy-ceae (red algae), over all from the families ofGigartinacieae and Solieriaceae. Carrageenan consists ofalternating 3-linked-b-D-galactopyranose and 4-linked-a-D-galactopiranose units. It precipitates as a gel in the pres-ence of cationic compounds: metal ions, amines, aminoacid derivates and water-miscible organic solvents (Tosaet al., 1979). Different isomeric forms of carrageenan canbe found, in function of the algal specific origin. Forinstance, i, j and k-carrageenan are primarily producedby Aghardhiella subulata and the gametophyte and tetrasp-orophyte of Chondrus crispus, respectively (Burdin andBird, 1994). Hardening processes in order to increase themechanical stability of carrageenan-based matrixes have


also been designed: Chamy et al. (1990) described a tech-nique of hardening for j-carrageenan immobilizing Sac-

charomyces cerevisiae by treatment with Al(NO3)3. Thistreatment revealed to increase cellular retention, a betterevacuation of gas from the reactor where immobilized cellswere cultured and an increase of 20% in ethanol productiv-ity by immobilized yeasts. Bacteria have also been success-fully immobilized (and stored during years) in carrageenanmatrixes (Cassidy et al., 1997). Travieso et al. (1996) com-pared the nutrient removal capacity of three microalgalspecies (C. vulgaris, Chlorella kessleri and Scenedesmus

quadricauda) immobilized in different matrixes (includingj-carrageenan hardened with 0.3 M KCl). But the stabilityof j-carrageenan beads was quite lower than that of Ca-alginate beads in the experimental conditions. The j-carra-geenan beads were partially destroyed after one week, thecells were liberated from the beads and a loss of bead struc-ture was reported after this time. Thus, the authors selectedCa-alginate instead of j-carrageenan for the describednutrient removal experiment.

Agar is a sulphated galactan obtained from some speciesof red algae (mainly from the genus Gelidium, Pterocladiaor Gracilaria) (Burdin and Bird, 1994). Agar is a thermo-reversible gel. The major gel forming component of agar(agarose) consists of a linear chain of sequences of (1–3)-linked-b-D-galactopyranosyl units and (1–4)-linkages to3,6-anhydro-a-D-galactopyranosyl units. Robinson et al.(1986), Codd (1987) and Papageorgiou (1987) cite thispolymer as suitable for immobilizing microalgal cells. Agarmatrixes for immobilizing living cells present a capital lim-itation: as it has been said, agar is a thermo-reversible gel.Dissolved at concentrations that will provide a goodmechanical structure, agar melts around 85 �C and solidi-fies between 35 and 40 �C. Species able to resist a shortthermal shock of this level should be selected for agarimmobilization. Attending to our experience, temperaturesover 30 �C could damage a wide variety of (at least) marinemicroalgae (except cyanophytes and some eukaryotic algaewell adapted to marshes and salt marshes environments,like some species from the genus Dunaliella, Nannochlorop-

sis or Tetraselmis). Agarose has been used as an immobili-zation matrix type fixed-bed for C. vulgaris in experimentsof Cu(II) biosorption (Aksu et al., 1998). In the agaroseparticle formation described by the authors, the agarosesolution containing C. vulgaris cells was dropped into edi-ble oil at 40 �C, although the temperature decreased afterdropping to 15 �C. The particles were then removed fromoily to aqueous phase by the addition of phosphate buffersolution. These authors reported that metal adsorptioncapacity of agarose and agarose–microorganisms systemwas lower than that reported for Ca-alginate in the sameconditions. Nevertheless, the presence of immobilized cellsincreased metal uptake capacity of both matrixes. Noefforts were made during this experiment in order to checkif immobilized cells were alive or not.

The most widely used polysaccharide gel for entrappingliving cells is alginate. Alginates constitute a family of

unbranched binary copolymers of 1–4-linked-b-D-mannu-ronic acid and a-L-guluronic acid in different proportionsand sequences (Smidsrød and Skjak-Braek, 1990), in func-tion of the organism and tissue they are isolated from.Commercial alginates are extracted from brown algae,mainly different species from the genus Laminaria (L.hyperborea, L. digitata, L. japonica) the species Macrocystis

pyrifera, Ascophyllum nodosum, Lesonia negrescens or spe-cies of the genus Sargassum, although all brown algae con-tain alginate in different proportions reaching up to 40% ofdry weight (Ertesvag and Valla, 1998). This substance has astructural function in those organisms. Some bacteria canalso produce alginates, as Azotobacter vinelandii (Smidsrødand Skjak-Braek, 1990) or Azotobacter chroococcum (Ert-esvag and Valla, 1998). Kidambi et al. (1995) describe theproduction of alginate by several phytopathogenic speciesof the genus Pseudomonas as a response to the additionof copper salts. The capacity of producing this substanceseems to be an evolutive strategy against the human useof copper-based phytosanitary products (high affinity ofalginates for copper will be further discussed in the text:Jang et al., 1995c), and additionally enhances bacterialability of adhesion to solid surfaces (Boyd and Chakra-barty, 1995).

A major advantage of alginate gel entrapment is thatimmobilized cells do not suffer extreme physical–chemicalcondition changes during the immobilization process. Per-meability, null toxicity and transparency of formed matriximply a very gentle environment for immobilized cells(Smidsrød and Skjak-Braek, 1990; Araujo and AndradeSantana, 1996). Alginates are used in industry as viscosifi-ers, stabilizers and gel-formers, film-formers or water-bind-ing agents (Ertesvag and Valla, 1998). The polymer issoluble in cold water and forms thermostable gels. Gelationof monovalent salts of this polysaccharide (normally Na-alginate) dissolved in water occurs when droplets of a mix-ture of cells (or enzymes) and alginate monovalent salts aremixed with a solution containing gel-forming ions. Gel for-mation is a very quick process. The most common cationused to form alginate gels is Ca2+. When the gel is intendedto be re-dissolved, media with sodium citrate (Hertzbergand Jensen, 1989) or phosphate (hexametaphosphate couldbe also used) can be used: calcium cations can be seques-tered by soluble anions or can be substituted in the matrixby monovalent cations in order to destabilize the structure.Na-alginate does not dissolve appropriately in sea or saltwater. Hertzberg and Jensen (1989) described a protocolfor immobilizing marine microalgae in alginate beads:Sodium chloride in adequate proportions for reaching sea-water salinity values and Na-alginate must be previously(separately) dissolved in distilled water and mixed after dis-solving. Marine microalgal cells can be then added to thesalty Na-alginate and dropped to the Ca2+ seawater solu-tion. These authors (Hertzberg and Jensen, 1989) immobi-lized seven marine microalgal species and checked thegrowth of entrapped cells. Pane et al. (1998) developed astudy of the viability of Tetraselmis suecica immobilized


in Ca-alginate beads and compared the cellular growth ofimmobilized and free cells, finding that the latter washigher, although chlorophyll content per cell in immobilizedcells was higher, possibly as a response to shading condi-tions provided by immobilization. Moreno-Garrido et al.(2005) checked the growth of immobilized cells and Ca-algi-nate bead stability in a 17-day experiment involving 11 mar-ine microalgal species, finding that the stability of beads canalso depend on the immobilized species. Attending to theresistance of the species to toxics, growth rates when immo-bilized and maintenance of the bead structure, thoseauthors selected T. suecica as a good target organism fortoxic accumulation experiments and P. tricornutum orChaetoceros gracilis as good target organisms for toxicitybioassays involving Ca-alginate immobilized microalgalcells. Loss of stability is a limitation for the use of Ca-algi-nate matrixes in sea, estuarine and brackish waters. The dis-solved monovalent cations (mainly sodium) in the mediacan substitute divalent cations and results in the loss ofstructure of matrixes. When shaking is applied, Ca-alginatebeads of 3–5 mm can dissolve in seawater in 24 h (unpub-lished data). In order to avoid these problems, some recentstrategies have been developed. Moreira et al. (2006) per-formed a study of alginate bead stability checking differentproportions of two types of alginates (isolated from M.

pyrifera and L. hyperborea) hardened with Ca2+ or Sr2+,immobilizing cells of P. tricornutum. Strontium divalentcations, as well as Ba2+ (Santos-Rosa et al., 1989), havebeen proposed as hardeners for alginates providing morestable gels in sodium and phosphate-enriched media. In astudy performed by Widerøe and Danielsen (2001), viabilityof entrapped cells in Ba, Ca and Sr-alginates was checkedon a human leukemic T cell line. Ba-alginate provided lowercellular growth rates, while cells exposed to calcium andstrontium grew in a similar way. In any case, inhibition ofgrowth of barium exposed cells was less than 20% in com-parison to the others. In support of the work described inMoreira et al. (2006), those authors confirmed that thebeads prepared using 4.9% of alginate from L. hyperborea

and hardened with a 4% strontium solution were found tobe the most stable and the most suitable for microalgalgrowth when they were exposed to natural field conditions.Small beads of Ca-alginate encapsulating bioactive sub-stances can be produced by emulsification and internal-ionotropic gelation (Poncelet et al., 1999). This method pro-poses dispersion of alginate and a calcium salt mixture (cit-rate) in canola oil by stirring. Then, certain volume ofcanola oil containing glacial acetic acid is added to theemulsion. After few minutes, this oil-bead suspension isadded to a calcium chloride solution in order to stabilizebeads and the oil is removed by washing with a surfactant.This technique has also been used for Ca-alginate encapsu-lating DNA (Quong et al., 1998) and it was improved byChan et al. (2002) in order to avoid possible clumping ofmicrospheres and excessive waste of calcium salts. Microen-capsulation has also been developed by co-use of alginateand polylysine (Jen et al., 1996). Other microencapsulation

strategy for microalgae is described by Joo et al. (2001): amixture of 2% sodium carboxymethyl cellulose, 2% CaCl2and microalgae is stirred in 0.8% sodium alginate till thecapsules are formed. After washing, the capsules are sub-merged in 2% CaCl2 for 20 min for the purpose of harden-ing. Those authors affirm that capsules had a betterresistance when compared with beads manufactured bythe ‘‘traditional’’ way (sodium salt dissolved and droppedin calcium solution), but do not report numerical data. Algi-nate is relatively cheap (overall when compared with otherimmobilizing matrixes), non-toxic, transparent enough, rel-atively stable and easy to use. As stability problems insodium or phosphate rich media have been avoided, thisimmobilization technique seems to be a very promising toolin microalgal biotechnology.

3. Effect of immobilization on microalgal cells

Toxicity of some immobilizing techniques on cells hasbeen discussed above (Blanco et al., 1999; Garbisu et al.,1991; Thepenier et al., 1985; Rangasayatorn et al., 2004;Urrutia et al., 1995). Overall, pre-polymers of syntheticfoams and resins use to be highly toxic for microalgae.When not so toxic immobilization techniques are applied,immobilized microalgal cells show, as minimum, a longerlag period when these are compared with free cells (Mal-lick, 2002; Vılchez et al., 1997). Some authors consider thissimilar to that occurring in free cultures (Lukavsky et al.,1986). After this period the specific growth rate (k) canbe very similar in free and immobilized cells (Lau et al.,1998; Mallick, 2002) although some reports confirmed thatit is lower than in the latter case: Pane et al. (1998) reporteda lag-phase of 5–6 days for immobilized T. suecica in Ca-alginate beads. It is common to find higher chlorophyllproduction in immobilized cells (Lau et al., 1998; Paneet al., 1998; Robinson et al., 1986). This is a phenomenonthat should be taken into account when estimations of bio-mass by chlorophyll are intended to be done. Hertzbergand Jensen (1989) demonstrated that the maximumamount of cells in culture could be higher for immobilizedalgae (P. tricornutum, in this case) than for free cells. Thetype of alginate and, overall, the initial cellular concentra-tion seem to be very important in maximum cellular den-sity that can be reached in immobilized cultures: thehigher initial cellular density, the higher final cellular den-sity to be reached in cultures. The latter phenomenon canbe explained because growing cells inside the immobilizingmatrix tend to form colonies. Maximum number of cellsper colony must be limited by some factors as nutrient dif-fusion or light (colonies are smaller in the inner part of thebeads than in the surface). Thus, the higher number of col-onies, the higher number of cells in the culture. In the samework, the authors found that S. costatum did not thrive sowell when immobilized: very high initial cell densities wereneeded to obtain stable cultures. Chaetoceros ceratosporumand Thalassiosira pseudonana also grew well in those exper-iments, while Emiliania huxleyi, Amphydinium carterae and


Scripsiella trochoidea did not grow when immobilized,although some of them appear to be metabolically activebecause during long periods of time these exhibited redautoflouorescence. Moreno-Garrido et al. (2005) per-formed a 17-day experiment on Ca-alginate beads immobi-lizing 11 different species belonging to eight differentmicroalgal taxonomic classes: Nannochloropsis gaditana

(Eustigmatophyceae); Heterocapsa sp. (Dinophyceae);Rhodomonas salina (Cryptophyceae); Isochrysis aff. galbana

(Prymnesiophyceae); T. pseudonana, C. gracilis, P. tricor-

nutum and S. costatum (Bacillariophyceae); Tetraselmis

chui (Prasinophyceae); P. cruentum (Rhodophyceae) andDunaliella salina (Chlorophyceae). Growth of cells and sta-bility of beads were checked during the performance of theexperiment. As Hertzberg and Jensen (1989) studied in thework cited before, Moreno-Garrido et al. (2005) found thatSkeletonema costaum did not grow when immobilized. Het-

erocapsa sp. also did not grow (as far as I know, there areno good results of immobilizing photosynthetic dinoflagel-lates in the literature), and N. gaditana resulted in unstablebeads in experimental conditions. The rest of the assayedspecies showed growth inside the beads, reaching sooneror later a stable number of cells (stationary equilibriumphase), except C. gracilis and I. galbana, which on day 17still showed continuous growth. Joo et al. (2001) immobi-lized four microalgal species (Chlorella minutissima, Pav-

lova lutheri, Haematococcus pluvialis and Dunaliella

bardawil) by two methods, and compared the growth ofimmobilized and free cells. The authors studied that cellsimmobilized by encapsulation by the use of 2% sodiumcarboxymethyl cellulose, as described before, grew betterfor the four species in bubble column bioreactors. Inhibi-tion, enhancement or no growth differences between freeand immobilized microalgal cells have been reported in dif-ferent works, such as Mallick (2002). Lukavsky et al.(1986) studied the morphology of immobilized cells in agarand Ca-alginate. Following those authors, which designedan experiment involving S. quadricauda and C. kessleri,cells suffered an initial lag phase (considered as comparableto that reported for free cells), followed by one to threedivisions. After that, cell division stops but giant cells inC. kessleri (which, following these authors, are commonin suspension exposed to sublethal conditions) are not usu-ally produced: cell shape and size do not show visiblechanges, organelles are recognizable and grains of starchappear inside the cells. In S. quadricauda, however, mon-strous cells use to appear. In any case, cells immobilizedin agar were less affected than those immobilized in algi-nate. For other immobilized microorganisms, manychanges in size and shape have been reported (Cassidyet al., 1996). Nevertheless, Hatanaka et al. (1999) did notfind differences in shape or chlorophyll content betweenimmobilized and free cells of Dunaliella parva. In general,alterations in shape of colonies are more frequent thanalterations in cell shapes (Mallick, 2002) when microalgalcells are immobilized. Vılchez et al. (1997) studied the via-bility of agar-immobilized Chlamydomonas reindhartii cells,

confirming that maximum growth was noticed near the sur-face of the beads. Diffusion of nutrients must also beimportant, apart from light, because no-photosyntheticorganisms such as the bacteria Nitrosomonas europea showthe same pattern of growth in carrageenan beads (Wijffelsand Tramper, 1989). Attending to Vılchez et al. (1997),these authors confirmed the presence of a thin skin of poly-mer surrounding colonies of C. reindhartii and attribute theconstant number of cells to the equilibrium between cellu-lar growth and release from the beads. The use of co-immo-bilized systems (Munoz and Guieysse, 2006) can improvethe growth of immobilized organisms, avoiding gas diffu-sive problems inside the immobilizing matrixes, as will bediscussed in Section 4.8.

Zeglinska (2005) (personal communication) reportedactive escape of filamentous cyanobacteria (Nodularia

spumigena) from alginate beads. Normally, motility of cellsis not strong enough to provide escape from beads, but inthe case of slithering filamentous cyanobacteria this move-ment seems to permit algae to escape from immobilizingmatrixes.

4. Current use of immobilized microalgal cells

Most frequent current uses of immobilized algal cells arethe culturing for metabolite production, improvement ofculture collections handling, obtaining of energy (via H2

or electricity power), nutrients, metal or organic pollutantremoval from aquatic media, measurement of toxicity andco-immobilization system production for different purposes.

4.1. Culturing for metabolite production

Microalgae has been widely used as biocatalysts in bio-transformation and de novo biosynthesis (Borowitzka andBorowitzka, 1988). Different types of bioreactors are ableto be used in microalgal biotechnology for different pur-poses (Mallick, 2002), immobilization techniques being aresponse to the problem of biomass losses in outflows(Nakasaki et al., 1989). Hatanaka et al. (1999) describeda reduction process mediated by Ca-alginate immobilizedD. parva, from hydroxyacetone to (R) 1,2-propanediol,which is a highly priced glycol. Production of propanediolby immobilized cells was found to be similar to that con-firmed for free cells. Tripathi et al. (2002) described bio-transformation of phenylpropanoid compounds toHPLC-detectable vanilla flavour metabolites by free andCa-alginate immobilized cells of H. pluvialis. Vanillin accu-mulation in immobilized cells is higher than in free cells (noexplanation of this phenomenon is given in the text, butreferences from other works with different microorganismsreviewed by those authors confirmed that it is not a rarecase). Transformation of nitrite into ammonium in a med-ium containing L-methionine-D,L-sulphoximine (MSX),and the inhibitor of glutamine synthetase, has beendescribed by Santos-Rosa et al. (1989) by the use of aBa-alginate immobilized mutant strain of C. reindhartii.


Maximum volumetric activity was 2700 lM h�1 in apacked-bed reactor. Ba-alginate was used in this casebecause a previous study demonstrated that cell leakagewas minor than that found in the same conditions forCa-alginate. Thepenier et al. (1985) studied production ofpolysaccharides by the unicellular red algae P. cruentumimmobilized in polyurethane foams. As it has been com-mented in Section 2, jelly colonies of microalgae could sur-vive to the toxicity of foam pre-polymers. At the beginningof the tests, no oxygen evolution was observed in immobi-lized cultures, possibly due to high degree of cellulardestruction, although after many flushing rinses, survivalcolonies began to grow again. When culture reaches sta-tionary phase, production of polysaccharides was notice-able (measured as an increase of medium viscosity).

4.2. Culture collection handling

Handling of a large microalgal culture collections is atedious, time-consuming task. Long-term storage is a veryinteresting topic related to microalgae culture handling,saving human and economic resources. Chen (2001)reported the maintenance of physiological activities of Ca-alginate entrapped S. quadricauda cells after three years ofstorage in darkness at 4 �C without culture medium, withthe sole starving symptom of pyrenoid disappearance. Afterreplacing long-term stored beads in fresh media, the num-ber of immobilized coenobia increased by near by 40 timesin four weeks (and pyrenoid was re-constructed). The sameauthor (Chen, 2003) repeated the experience with I. galbana,achieving good results after one year of storage. Hertzbergand Jensen (1989) stored Ca-alginate cultures of P. tricornu-

tum during one year in diminished light at 4 �C and con-firmed immediate production of oxygen in a Clark-typeelectrode when beads were transferred to light. Lukavsky(1988) immobilized 31 strains of cyanophytes and eukary-otic algae in 2% agar (inside the warm agar, not seeded onthe surface), and kept cultures at reduced temperature(10 �C). After 32 months most of the strains were still met-abolically active. Joo et al. (2001) immobilized four micro-algal species (D. bardawil, C. minutissima, P. lutheri and H.

pluvialis) in Ca-alginate capsules and beads in order to gethighly dense cultures. Those authors achieved higher con-centration of encapsulated cells than in the case of cultureswith free cells or bead-type immobilized cells. Response ofD. bardawil and H. pluvialis in bubble column reactorswas specially high, reaching values five times higher thanthat for free cells. Romo and Perez-Martınez (1997) alsostored Pseudanabaena galeata in Ca-alginate beads for 14–18 months. Recuperation of cells in fresh cultures was notdifferent to that checked for standard cultures. Hertzbergand Jensen (1989) also pointed out that immobilization isan interesting way to produce colonies from disperse cul-tures. Extending their affirmation, the technique could alsobe used as a way of producing clonal colonies from naturalmicroalgal assemblages in order to isolate scarce cells of adeterminate species via posterior micromanipulation. Novel

techniques have been developed in order to facilitate culti-vation of algae. Nowak et al. (2005) designed a systembased in a 96-well plate where a membrane filter, whichimmobilizes algal strains, constitutes the bottom of eachwell. This system allows culturing of several microalgalstrains with less effort, time and money.

4.3. Obtaining energy (electricity or hydrogen)

It is said that hydrogen is the fuel of the future due to itshigh conversion efficiency, recyclability and non-pollutingnature (Das and Veziroglu, 2001). Bioproduction of hydro-gen has demonstrated to be environmental friendly and lessenergy consumer when compared with thermochemical andelectrochemical processes (Kapdan and Kargi, 2006). Somealgae are able to produce hydrogen in stress conditions(Melis, 2002) such as the deprivation of sulphur-nutrientsin green algae, which causes a reversible inhibition of theoxygenic photosynthetic processes. Lack of sulphur pre-vents the protein synthesis and algae cannot perform theturnover of specific photosystem-II reaction centre protein.Then, photochemical activity of PSII declines in theabsence of sulphur and oxygen is released to the media atlower rates than O2 consumption, achieving anaerobicalgal cultures in the dark if they are sealed and sulphur-deprived. The same occurs when O2 is directly removedfrom the media. Some experiments related to this topichave recently been developed on C. reindhartii and otherspecies (Dante, 2005; Laurinavichene et al., 2006; Markovet al., 2006; Polle et al., 2002). C. reindhartii seems to be apromising organism for further studies in this field. Co-fir-ing of microalgal biomass with coal is another power-gen-erating procedure. CO2 produced in power-plant fuel gascan be used to improve microalgal growth. Microalgal bio-mass obtained in this way is burned again in the samepower plant (Kadam, 2002). This could be a suitable wayof reducing the high amounts of carbon dioxide (around26 Gt per year, following the same author) produced byhumans and helps the high CO2 producing countries (TheUnited States generates 5.7 Gt, it means about 22% ofworldwide anthropogenic emissions) to reach CO2 levelsin agreement with the Kyoto Protocol (van Vuuren et al.,2006). In any case, molecular hydrogen production seemsto be the target in microalgal-based energy production, asmedium and long-term endurance of thermal electric plantsbased in firing materials should be severely studied by envi-ronmentalists and governments, due to the recognized pol-lutant capacity of those types of plants. Some efforts hadalso been made during the eighties in order to generateelectricity by photovoltaic cells containing photosyntheticbiocatalysts, as reviewed by Papageorgiou (1987), but norecent references have been found with respect to this topic.

4.4. Nutrient removal

Cultivation of algae in wastewater containing nutrientsoffers the combined advantages of treating water and pro-


duction of algal biomass, which can be industriallyexploited (Mallick, 2002). A limitation in these processesis the high price and time-consumption of centrifugationor filtration. Immobilization techniques have tried to solvethese problems (Proulx and de la Noue, 1988; Tam andWong, 2000; Travieso et al., 1996; Vılchez and Vega,1994, 1995; Vılchez et al., 2001). Immobilized P. laminosum

in polyurethane and polyvinyl foam has been used in con-tinuous flow reactors designed in order to remove nitratefrom water (Garbisu et al., 1991). Comparison was donebetween adsorbed and entrapped cells. As it has been said,pre-polymers of polyurethane foams are toxic to cells.Those authors concluded that entrapment was not a suit-able method for immobilizing living cells in those matrixes,and experiments were only performed on adsorbed cells. Inthe work described, only nitrate was removed from themedium by adsorbed cells in light, and nitrite or ammo-nium was not released (less than 1 mg L�1 of both nutrientsin all the samples). Using chlorophyll as an indicator ofmicroalgal biomass (this can lead to errors if a correlationbetween chlorophyll and number of cells is done based onfree cells, as it has been commented), immobilized P. lami-nosum removed nitrate at 1.1 lM mg�1 chl h�1, while freecells removed 2.6 lM mg�1 chl h�1 in batch cultures. Inreactors (no free cells can be used here), removal of nitratehad an efficiency of 90% during three months. The authorscomment that a packed bed is a more adequate system thana fluidized reactor, because in the latter tumbling and col-lisions between particles result in desorption of cells. Atlow concentrations, consumption of nutrients by immobi-lized microalgal cells is limited, possibly due to limitationin nutrient diffusion through immobilizing matrixes. Thus,Garbayo et al. (2000) found that Ca-alginate immobilizedC. reindhartii cells did not consume nitrate when concen-tration of this nutrient was below 0.14 mM, while free cellsalmost fully consumed it. The authors postulated thatnitrate consumption inhibition could occur due to the pres-ence of nitrite. Mallick (2002) reviewed some works statingthat exponentially growing microalgal cells removed morenutrients (nitrate and phosphate) than aged cultures.Strains of Scenedesmus intermedius and Nannochloris sp.were isolated from different sources of pig manure in orderto design a depuration system for the removal of macronu-trients from farm wastewaters (Jimenez-Perez et al., 2004).Nitrogen and phosphorus uptake from free and Ca-algi-nate immobilized cells were compared in this work, findingto be slightly lower in immobilized systems (but in the sameorder of magnitude). Nevertheless, the uptake of nutrientsby those autochthonous species was markedly higher thanthat measured for common commercial species, probablydue to a better adaptation to high nutrient concentrations.

In seawater selection of resistant taxons for depurationassays is also an important topic. Thakur and Kumar(1999) selected D. salina in order to perform nutrient(ammonium, nitrate and phosphate) uptake experiments.This alga is halotolerant and could be used in waters of awide range of salinities. In this concrete experiment, immo-

bilized cells always removed more nutrients than free cells.After 36 h, the levels of removed nitrate, ammonium andphosphate were 62%, 42% and 65% of initial concentra-tions, respectively. Dunaliella species, thus, seems to be agood target organism for accumulation assays in watersof variable salinity (Donmez and Aksu, 2002). Immobiliza-tion techniques improve cost-effective phycoremediationprocesses (Olguın, 2003). The removal of nutrients fromanimal wastewaters is a very interesting topic as manyunderground water bodies in several regions of the worldare threatened of suffering eutrophication (Travieso Cor-doba et al., 1995a,b). Recently, co-immobilized systemshave been developed in order to enhance nutrient removalfrom wastewaters (see Section 4.8).

4.5. Metal removal

Vegetal biomass (Spinti et al., 1995) and especiallymacro- (Valdman et al., 2001; Al-Rub et al., 2004) and mic-roalgae are known to accumulate high amounts of metalfrom their environment (Stark and Rayson, 2000; Traviesoet al., 2002). This capacity has been exploited for differentpurposes (Aksu and Acikel, 1999; Burdin and Bird, 1994;Greene and Bedell, 1990; Hashim et al., 2000). Silica immo-bilized-algal biomass (Pilayella littoralis) has been pro-posed to be used as a biosorbent for metal pre-concentration before measuring in analytical devices(inductively coupled plasma optical emission spectrometry)(Carrilho et al., 2003). But the main purpose of the use ofimmobilizing algae binding metals has been detoxificationand metal recovery (Greene and Bedell, 1990). The latteris possible because a great part of the metal bound to cel-lular surfaces and immobilizing systems is able to be des-orbed via acid treatment. Other resins, such asAmberlite�, have been used in order to immobilize micro-organisms set aside for metal removal (Baytak and Turker,2005). Sorption of metals on algae seems to be not only asimple adsorption process, but also an exchange of ions,where Ca2+ is often replaced (Crist et al., 1994). Bindingof metals to algal surface occurs in living and non-livingalgae (Greene and Bedell, 1990), cell surface area being amajor parameter in the uptake of metals by microalgae(Khoshmanesh et al., 1997). Dead cells can be very efficientin accumulating metals (Donmez et al., 1999). Blanco et al.(1999) used biomass of the cyanobacterium P. laminosum

immobilized in polysulphone and epoxy resin beads forCu(II), Fe(II), Ni(II) and Zn(II) sequestering. They foundthat the amount of biosorbed metal increased with biomassand the amount of metal available. Biosorption–desorption(acid-mediated) cycles with this immobilized system main-tain the efficiency after at least 10 cycles. Copper is selec-tively adsorbed by alginates (Alhakawati and Banks,2004; Jang et al., 1995b,c; Nestle and Kimmich, 1996). Janget al. (1995a) considered the possibility of adding copper-sequestering agents to alginate gels in order to enhancecopper recovery from the media. Addition of sodium poly-styrenesulphonate (NaPSS) to the sodium alginate resulted


in an increase of copper adsorption, but the addition ofcells of the cyanophyte Microcystis sp. resulted in an evenbetter result. Accumulation of metals by immobilizing sys-tems containing microalgae seems to present two phases(Malik, 2004). Garnham et al. (1992a) developed an exper-iment involving immobilized Chlorella salina in Ca-alginatebeads, accumulating radioisotopes of metals (60Co, 54Mnand 65Zn). The authors found a rapid biosorption non-dependent on light, temperature or metabolic inhibitor(carbonylcyanide n-chlorohydrazone: CCCP), followed bya slower accumulation phase depending on cellular metab-olism. Similar results have been found by Moreno-Garridoet al. (1998, 2002): Part of the metal accumulated cannot bewashed by solutions containing high concentrations of che-lating EDTA. It is supposed that this metal is absorbedinto the cells by active or passive methods (Greene andBedell, 1990) and not adsorbed to the cell surface or theimmobilizing matrix. Packed-bed columns containingimmobilized cells seem to be very efficient in the removalof metals from aquatic media (more than fluidized-bed orair-lift reactors, Moreno-Garrido et al., 2002). Aksu et al.(1998) studied biosorption of copper(II) in Ca-alginateand agarose immobilized C. vulgaris. Those authors didnot find a significant increase in metal adsorption due tothe presence of immobilized algae. But Moreno-Garridoet al. (2002) found great differences in Ca-alginate beadssystem immobilizing cells of N. gaditana when comparedwith systems without immobilized cells: Beads containingcells accumulate practically all Cu in media (as free cellsdid) and 80% of Zn, while beads without cells accumulatednear by 80% of Cu but did not accumulate measurableamounts of Zn. Similar results can be found when deadcells were employed: no Zn was removed when dead cellsare immobilized in Ca-alginate beads.

Isolating microalgal strains from polluted waters is asuitable tool for selecting metal-tolerant and highly accu-mulating cells. Khattar et al. (1999) isolated a strain ofAnacystis nidulans from polluted sites. This strain was ableto grow in a medium up to 100 lM of chromium, beingable to accumulate 43 nM of Cr per mg of microalgal pro-tein (as estimative of biomass) immobilized in agar (freecells were able to accumulate 35 nM of Cr) in the experi-mental conditions. Akhtar et al. (2004) reported recoveryof nickel from electroplating industrial effluents by theuse of C. sorokiniana immobilized in loofa sponges by nat-ural adsorption. These authors achieved a maximum C.sorokiniana biomass of 261 ± 22 mg g�1 of dry spongeafter 24 days of incubation. Adsorption was always higherin immobilized systems than that measured for free cells(loofa sponges without immobilized cells adsorbed a smallamount of this metal). After adsorption, addition of acidssuch as HCl and H2SO4 resulted in liberation of more than99% of adsorbed metal. Moreno-Garrido et al. (2005) usedTetraselmis chuii, a very stress-resistant taxon, as an immo-bilized organism to remove Cu and Cd from marine waters.After 24 h exposition, all Cu and 20% Cd were removedfrom the media by Ca-alginate immobilized populations

of T. chuii. Rangasayatorn et al. (2004) found a maximumCd accumulation capacity of 70.9 mg g�1 biomass forimmobilized cells in Ca-alginate matrixes. Cells were suc-cessfully used during five adsorption–desorption consecu-tive processes. During these cycles, system adsorptioncapacity was reduced from around 94% to near by 66%of total cadmium. Microalgal living cells used to accumu-late higher levels of metals than dead microalgal biomass(Moreno-Garrido et al., 1998). First, divalent cationsbecame less soluble at high pH values. Active (photosyn-thetic) cells provide a high-pH environment in the immedi-ate surface cellular layer, increasing thus adsorption to thecellular surface. Second, there are processes of absorption(dependent on cellular metabolism) of metals by microal-gae (Garnham et al., 1992a,b; Moreno-Garrido et al.,2002), slower than the adsorption process that increasesthe capacity of living algae to accumulate those pollutants.

Noble metals can also be accumulated by algae andselectively eluted from them (Guo et al., 2000). Gee andDudeney (1987) described the adsorption of gold from adissolved metal mixture by C. vulgaris and S. platensis

immobilized in Ca-alginate. In this case, selective desorp-tion of Fe and Au was performed by acidic pH adjustmentand acidic thiourea addition, respectively. Guo et al. (2000)also described the uptake of neodymium, a rare lantha-noid, by living and fossil (570 million years old) algae.

4.6. Organic pollutants removal

Aksu (2005) made a review about the biosorption oforganic pollutants and found only few data on microalgae.In the same year, Aksu and Tezer (2005) studied the bio-sorption of three reactive dyes onto dead biomass of C. vul-

garis, finding that dye sorption was highly dependent onpH: the optimum pH value for adsorption was 2.0. Temper-ature also affected the process in a proportional inverseway. Oh et al. (2000) immobilized yeast cells in polyure-thane foams in order to absorb and degrade oil on watersurface. Few works describing algal capacity for degradingoil have been published. A recent paper from Chaillan et al.(2006) described the appearance of cyanobacterial mats(Phormidium animale) in petroleum-polluted site. They con-cluded that there is no evidence of biodegradation of crudeoil by cyanophytes, but from other organisms present in themat formed by Phormidium involved in degradation pro-cesses, Biofilm-forming bacteria covering macroalgae canalso be able to degrade oil (Radwan et al., 2002).

C. sorokiniana in aggregation with bacteria was able tosuccessfully remove salicylate from a photobioreactor inan experiment described by Munoz et al. (2006). The roleof each organism in the biosystem is not clear enough,but degradation seems to be performed by the bacterialpart of the symbiosis system. Normally, organic pollutantsare more easily degraded by bacteria than by algae (Prietoet al., 2002). Hydrocarbons, nevertheless, have beenreported to be degraded by Ca-alginate immobilized color-less P. zopfii (Suzuki et al., 1998; Yamaguchi et al., 1999).


In free and immobilized systems, 100% of a mixture oftetradecane, pentadecane and hexadecane (0.1% (v/v) eachone) was degraded. Special care in the experimental designwas taken in order to ensure that evaporation was not thecause of the elimination of hydrocarbons from the media.Every 14 days, cells were washed and re-used, resulting inidentical response from immobilized cells. Other cases oftrophic use of organic pollutants seem to be the Chryso-phyte Ocrhomonas danica (Semple and Cain, 1996; Semple,1998), which is able to heterotrophically grow on phenoland phenolic mixtures. No reports about immobilizationof this species have been found.

4.7. Measuring toxicity

Bulk chemical characterization of waters and sedimentsdo not provide information about toxicity (Munawar andMunawar, 1987), except in cases of extreme pollution.Thus, bioassays have been designed in order to detect tox-icity of effluents, sediments or substances on organisms.Microalgae have been found to be sensitive organisms todifferent pollutants (Leon et al., 2001; Radix et al., 2000)in toxicity bioassays (Bitton and Dutka, 1986), possiblydue to their high surface/volume ratio. Their key role infreshwater and marine aquatic trophic nets implies thenecessity of developing suitable toxicity tests in order tocount with efficient tools to be used by researchers andauthorities when it is required. In situ experiments havebeen designed in order to increase environmental relevanceof toxicity tests (Moreira dos Santos et al., 2002, 2004;Moreira et al., 2006; Munawar and Munawar, 1987), asmanipulation of samples when carried to the laboratoryis then avoided and natural conditions of light, tempera-ture or pH fluctuations are maintained. Bozeman et al.(1989), in a pioneering work, compared the toxicity ofseven pollutants of different origin (Cd, Cu, Glyphosate,Hydrothol, Paraquat, pentachlorophenol and sodiumdodecyl sulphate) to free and immobilized cells of the greenmicroalga Selenastrum capricornutum (currently Pseud-okirchneriella subcapitata), suggesting the possibility ofthe use of immobilized systems in in situ toxicity experi-ments. Following those authors, differences in toxicity forfree and immobilized algae varied from no significant dif-ferences for copper and pentachlorophenol to nearby fourtimes more sensitive for free cells in the case of Glyphosateor Paraquat. Admiraal et al. (1999) performed an experi-ment on sand and natural glass-attached microbenticassemblages of algae and bacteria in a metal pollutedstream in the river Dommel (Belgium). The authors com-pared the sensitivity of those assemblages to zinc, findingdifferent sensitivities in function of the origin of the assem-blages (the most polluted origin, the lower sensitivity). Pro-tection against toxicity in immobilized cells is reported indifferent works (Cassidy et al., 1996). Awasthi and Rai(2005) demonstrated lower inhibition of nitrate uptake inS. quadricauda immobilized (with respect to free cells) whenexposed to Ni, Zn or Cd. In this work, no metal measure-

ments were performed in the media, and the easiest expla-nation is the removal of part of the metals by theentrapping matrix, being thus less available for cells. Butremoval of toxicants by immobilizing matrixes would notexplain all cases of less toxicity of immobilized cells. Sur-factants are not so selectively adsorbed by Ca-alginates.Moreno-Garrido et al. (2007) found less toxicity for immo-bilized cells of P. tricornutum exposed to sediments spikedwith surfactant lineal alkylbenzene sulphonate (LAS) thanfor free cells. Lower diffusivity of toxicants in beads (Jang,1993) should also explain, at least in part, lower toxicity toentrapped cells. This species (P. tricornutum) seems to be agood microalgal species to be used in toxicity tests involv-ing immobilized algae. It is one of the species suggested byISO (1995) to test water quality by the use of growth inhi-bition tests: it is cosmopolitan, with low nutrient require-ments, fast growth and good sensitivity to toxicants.Additionally, it has been used by several authors in freeand immobilized toxicity tests (Cid et al., 1995; Kos-akowska et al., 2004; Mayasich et al., 1986; Moreiraet al., 2006; Moreira dos Santos et al., 2002; Morelli andPratesi, 1997; Morelli and Scarano, 2001; Moreno-Garridoet al., 2007; Overnell, 1975; Pavlic et al., 2005; Wiegmanet al., 2002). Immobilization by gel entrapment is also avery interesting topic for in situ microalgal experimentdesign because it provides protection to microalgae in frontof grazers (Cassidy et al., 1996; Faafeng et al., 1994). Mic-roalgal grazers cannot be easily eliminated from waters orsediments due to the small size that those organisms canachieve: nematodes and, over all, amoebas or ciliates canpredate on microalgal cells slightly smaller than them.Twist et al. (1997) developed a method of in situ biomoni-toring using Ca-alginate immobilized Scenedesmus subspic-

atus for the assessment of eutrophication in surface waters.A clear advantage of this technique is that local flora canbe isolated and incorporated to the biomonitor. Of course,when Ca-alginate is used in natural (or micro and meso-cosms simulating natural) environment, limitation of thetechnique is degradation of beads. In freshwater streamsthis limitation seems to be a couple of weeks. In marineenvironments (Faafeng et al., 1994), duration of beads isquite shorter (a few days) (Moreira dos Santos et al.,2002). Improvement in bead fabrication is trying to avoidthose problems (Moreira et al., 2006). Periphytic algae nat-urally established on glass slides have been used in in situ

toxicity tests conducted to test the impact of heavy metalsresuspended by dredging works on microalgae (Nayaret al., 2003). High concentrations of Zn (up to17,240 mg L�1), Cu (up to 11 mg L�1) and Cd (up to1.8 mg L�1) were found, as a result of dredging, in theaqueous phase, reducing periphyton biomass between95% and 100% for polluted waters.

Biosensors of different types involving microalgal cellshave been designed in order to detect pollutants in the envi-ronment. Lukavsky and Marsalek (1997) immobilized S.

capricornutum in 2% agar in order to use it as a biosensorfor Cr6+ toxicity. Sensitivity of this biosensor was the same


found for other bioassays, including growth inhibition test.Frense et al. (1998) used S. subspicatus immobilized on afilter paper and covered with alginate in an optical biosen-sor based in the chlorophyll a fluorescence as biomarkerfor pollutants in water and soil extracts, as a pre-selectionprotocol before sending samples to high-cost standardanalysis. C. vulgaris has also been used in optic biosensorsin order to determine the toxicity of herbicides (Naessenset al., 2000) such as atrazine, simazine and diuron, com-monly used in cereal cultures. In this case, algal immobili-zation was performed on GF/C Whatman filters. Filterpaper disks immobilizing algae have also been used bySanders et al. (2001) in biosensors designed in order todetect chemical warfare agents. Not optic, but ampero-metic algal-based sensors have also been designed (Shit-anda et al., 2005), taking advantage of variations in thephotosynthetically produced oxygen. Limitations of toxic-ity testing using free or immobilizing microalgae arerestricted to those toxicants that affect structures presentin the algal cells. It means that pollutants affecting bonedevelopment or nervous system will not be easily detectedwith microalgal-based bioassays. In contrast, toxicantsaffecting photosynthesis (as copper ions or herbicides) willbe more appropriately detected with vegetal cells (such asmicroalgae) bioassays. Adequate aquatic toxicity studiesshould cover organisms from different levels, but microal-gae should never be forgotten due its basal position inthe trophic chain.

4.8. Co-immobilized systems

Recent efforts have been made in the field of co-immobi-lization (Nagase et al., 2006). De-Bashan et al. (2002a,b,2004) co-immobilized Chlorella with a microalgaegrowth-promoting bacterium (Azospirillum brasiliense) inCa-alginate beads. This bacterium is not able to removenutrients from wastewaters, but enhances growth of immo-bilized algae. Co-immobilized biological system removedhigher percentages of nutrients from wastewaters (100%of ammonium, 15% nitrate and 36% of phosphorus) whencompared with immobilized algal cells without bacteria(75% ammonium, 6% nitrate and 19% phosphorus).Munoz and Guieysse (2006) reviewed the interactionsbetween algae and bacteria in processes designed for thetreatment of hazardous contaminants. Production of oxy-gen by algae improves degradation of substances that mustbe degraded aerobically. Both bacteria and algae couldproduce defending substances against the other co-immobi-lized organism. Increase of pH values due to photosynthe-sis and increase of oxygen in the media could also slowdown bacterial growth when co-immobilized with algae.On the other hand, consumption of CO2 and extracellularmatter production (such as exopolysaccharydes) by algaecan enhance bacterial growth rate, as well as CO2 andgrowth promoter substances production by bacteria canenhance microalgal growth. The same authors recognizedthat most widely used immobilization techniques involve

weak and expensive matrices (when used in large scale),and propose other approaches such as photobioreactorswith the bacterial–algal microcosm attached on the reactorwalls.

5. Future perspectives

Immobilization techniques can be used in molecularbiology, as plasmid stability in immobilized cells has beenreported (Cassidy et al., 1996). Risk of unwanted muta-tions is reduced when cells are immobilized (Codd, 1987).Genetic manipulation of immobilized cyanobacteria(hydrogenase negative gene) could also improve hydrogengeneration processes (Das and Veziroglu, 2001). Immobi-lized microalgae have been recently used as a tool for waterquality control in fish culture. Chen (2001) studied thatammonium concentrations decreased in tilapia culturescontaining immobilized cells of S. quadricauda (freshwatercultures) and clam cultures (Chen, 2003) containing I. gal-

bana (seawater cultures). In the latter case, slow liberationof cells from immobilizing beads provided a continuousinput of food for filtering clams. This could reduce costsof clam culturing when compared with traditionalmethods.

The use of microalgae in the design of biosensors is avery recent and interesting topic in biotechnology. Chou-teau et al. (2004) designed a conductometric biosensor withC. vulgaris cells (the authors pointed out that the use ofwhole cells is quite cheaper than that of isolated enzymes)in order to measure the inhibition of alkaline phosphataseactivity as a bio-indicator of toxic stress. Volatile com-pounds (such as formaldehyde) have also been detectedby the use of multiple-strain algal sensor chips designedby Podola et al. (2004). Genetically modified organismscan also be used in sensors, such as the strain of Synecho-

coccus employed by Schreiter et al. (2001): this organismharbours a gene encoding for the luciferase from Vibrio

harveyi under the control of an inducible alkaline phospha-tase promoter, which can be induced by phosphorus limita-tion. The resultant ‘‘CyanoSensor’’ is able to detect 0.3–8 lM of PO3�

4 and respond to other organic phosphorussources. This sensor is also storable for three weeks at 4 �C.

Combinations of solar energy and algal immobilizationtechnologies can be successfully used in industrial processes(Mallick, 2002). In the same way, studies about productionof energy via photosynthetically generated H2 are a recent,promising field of research that has been reduced to date togreen freshwater algae. Other taxonomic groups should betested in order to optimize molecular hydrogen production.

Acknowledgements

This work has been granted by the Spanish NationalPlan of Science and Technology Research, under the pro-jects ‘‘Use of immobilized systems for marine microalgaein the incorporation and evaluation of toxic substances inmarine ecosystems’’ (REN2001-2095/MAR) and ‘‘A new


ecologically relevant tool in environmental toxicology:microphytobenthos for the environmental quality assess-ment of estuarine and coastal sediments (MECASEC)’’(CTM2006-01473/MAR).

References

Admiraal, W., Blanck, H., Buckert-de Jong, M., Guasch, H., Ivorra, N.,Lehmann, V., Nystrom, B.A.H., Paulsson, M., Sabater, S., 1999.Short-term toxicity of zinc to microbenthic algae and bacteria in ametal polluted stream. Water Res. 33, 1989–1996.

Ahmadi, M., Vahabzadeh, F., Bonakdarpour, B., Mehranian, M., 2006.Empirical modeling of olive oil mill wastewater treatment using loofa-immobilized Phanaerochaete crysosporium. Process Biochem. 41, 1148–1154.

Akhtar, N., Iqbal, J., Iqbal, M., 2004. Removal and recovery of nickel(II)from aqueous solution by loofa sponge immobilized biomass ofChlorella sorokiniana: characterization studies. J. Hazard. Mater. B108, 85–94.

Aksu, Z., 2005. Application of biosorption for the removal of organicpollutants: a review. Process Biochem. 40, 997–1026.

Aksu, Z., Acikel, U., 1999. A single-staged bioseparation process forsimultaneous removal of copper(II) and chromium(VI) by using C.

vulgaris. Process Biochem. 34, 589–599.Aksu, Z., Egrtli, G., Kutsal, T., 1998. A comparative study of copper(II)

biosorption on Ca-alginate, agarose and immobilized C. vulgaris in apacked-bed column. Process Biochem. 33, 393–400.

Aksu, Z., Tezer, S., 2005. Biosorption of reactive dyes on the green algaChlorella vulgaris. Process Biochem. 40, 1347–1361.

Alhakawati, M.S., Banks, C.J., 2004. Removal of copper from aqueoussolution by Ascophyllum nodosum immobilised in hydrophilic poly-urethane foam. J. Environ. Manage. 72, 195–204.

Al-Rub, F.A.A., El-Naas, M.H., Benyahia, F., Ashour, I., 2004.Biosorption of nickel on blank alginate beads, free and immobilizedalgal cells. Process Biochem. 39, 1767–1773.

Araujo, A.A., Andrade Santana, M.H., 1996. Aerobic immobilized cells inalginate gel particles of variable density. Appl. Biochem. Biotech. (57/58), 543–550.

Archambault, J., Volesky, B., Kurz, W.G.W., 1990. Development ofbioreactors for the culture of surface immobilized plant cells.Biotechnol. Bioeng. 35, 702–711.

Awasthi, M., Rai, L.C., 2005. Toxicity of nickel, zinc, and cadmium tonitrate uptake in free and immobilized cells of Scenedesmus quadric-

auda. Ecotox. Environ. Safe 61, 268–272.Baytak, S., Turker, A.R., 2005. The use of Agrobacterium tumefaciens

immobilized on Amberlite XAD-4 as a new biosorbent for the columnpreconcentration of iron(III), cobalt(II), manganese(II) and chro-mium(III). Talanta 65, 938–945.

Bitton, G., Dutka, B.J. (Eds.), 1986. Toxicity Test Using Microorganisms.CRC Press, Boca Raton, FL.

Blanco, A., Sanz, B., Llama, M.J., Serra, J.L., 1999. Biosorption of heavymetals to immobilized Phormidium laminosum biomass. J. Biotechnol.69, 227–240.

Borowitzka, M.A., Borowitzka, J.L., 1988. Micro-algal biotechnology.Cambridge University Press, Cambridge.

Boyd, A., Chakrabarty, A.M., 1995. Pseudomonas aeruginosa biofilms:role of the alginate exopolysaccharide. J. Ind. Microbiol. 15, 162–168.

Bozeman, J., Koopman, B., Bitton, G., 1989. Toxicity testing usingimmobilized microalgae. Aquat. Toxicol. 14, 345–352.

Brandini, F.P., da Silva, E.T., Pellizari, F.M., Fonseca, A.L.O., Fernan-des, L.F., 2001. Production and biomass accumulation of periphyticdiatoms growing on glass slides during a 1-year cycle in a subtropicalestuarine environment (Bay of Paranagua, southern Brazil). Mar. Biol.138, 163–171.

Burdin, K.S., Bird, K.T., 1994. Heavy metal accumulation by carrageenanand agar producing algae. Bot. Mar. 37, 467–470.

Carrilho, E.N.V.M., Nobrega, J.A., Gilbert, T.R., 2003. The use of silica-immobilized brown alga (Pilayella littoralis) for metal preconcentra-tion and determination by inductively coupled plasma optical emissionspectrometry. Talanta 60, 1131–1140.

Cassidy, M.B., Lee, H., Trevors, J.T., 1996. Environmental applications ofimmobilized cells: a review. J. Ind. Microbiol. 16, 79–101.

Cassidy, M.B., Lee, H., Trevors, J.T., 1997. Survival and activity of lac-lux marked Pseudomonas aeruginosa UG2Lr cells encapsulated in j-carrageenan over four years at 4 �C. J. Microbiol. Methods 30, 167–170.

Chaillan, F., Gugger, M., Saliot, A., Coute, A., Oudot, J., 2006. Role ofcyanobacteria in the biodegradation of crude oil by a tropicalcyanobacterial mat. Chemosphere 62, 1574–1582.

Chamy, R., Nunez, M.J., Lema, J.M., 1990. Optimization of thehardening treatment of S. cerevisiae bioparticles. Enzyme Microb.Tech. 12, 749–754.

Chan, L.W., Lee, H.Y., Heng, P.W.S., 2002. Production of alginatemicrospheres by internal gelation using an emulsification method. Int.J. Pharm. 242, 259–262.

Chen, Y.-C., 2001. Immobilized microalga Scenedesmus quadricauda

(Chlorophyta, Chlorococcales) for long-term storage and applicationfor water quality control in fish culture. Aquaculture 195, 71–80.

Chen, Y.-C., 2003. Immobilized Isochrysis galbana (Haptophyta) for long-term storage and applications for feed and water quality control inclam (Meretrix lusoria) cultures. J. Appl. Phycol. 15, 439–444.

Chouteau, C., Dzyadevych, S., Chovelon, J.M., Durrieu, C., 2004.Development of novel conductometric biosensors based on immobi-lized whole cell Chlorella vulgaris microalgae. Biosen. Bioelectron. 19,1089–1096.

Cid, A., Herrero, C., Torres, E., Abalde, J., 1995. Copper toxicity on themarine microalga Phaeodactylum tricornutum: effects on photosynthe-sis and related parameters. Aquat. Toxicol. 31, 165–174.

Codd, G.A., 1987. Immobilized micro-algae and cyanobacteria. Br.Phycol. Soc. Newslett. 24, 1–5.

Crist, R.H., Martin, J.R., Carr, D., Watson, J.R., Clarke, H.J., Crist,D.R., 1994. Interaction of metals and protons with algae. 4. Ionexchange vs. adsorption models and reassessment of scatchard plots;ion-exchange rates and equilibria compared with calcium alginate.Environ. Sci. Technol. 28, 1859–1866.

Danilov, R.A., Ekelund, N.G.A., 2001. Comparison of usefulness of threetypes of artificial substrata (glass, wood and plastic) when studyingsettlement patterns of periphyton in lakes of different trophic status. J.Microbiol. Methods 45, 167–170.

Dante, R.C., 2005. Hypotheses for direct PEM fuel cells applications ofphotobioproduced hydrogen by Chlamydomonas reindhartii. Int. J.Hydrogen Energy 30, 421–424.

Das, D., Veziroglu, T.N., 2001. Hydrogen production by biologicalprocesses: a survey of literature. Int. J. Hydrogen Energy 26,13–28.

De-Bashan, L.E., Bashan, Y., Moreno, M., Lebsky, V.K., Bustillos, J.J.,2002a. Increased pigment and lipid content, lipid variety, and cellspopulation size of the microalgae Chlorella sp. when co-immobilized inalginate beads with the microalgae-growth-promoting bacteriumAzospirillum brasilense. Can. J. Microbiol. 48, 514–521.

De-Bashan, L.E., Moreno, M., Hernandez, J.P., Bashan, Y., 2002b.Removal of ammonium and phosphorus ions from synthetic waste-water by the microalgae Chlorella vulgaris coimmobilized in alginatebeads with the microalgae growth-promoting bacterium Azospirillum

brasiliense. Water Res. 36, 2941–2948.De-Bashan, L.E., Hernandez, J.P., Morey, T., Bashan, Y., 2004. Micro-

algae growth-promoting bacteria as ‘‘helpers’’ for microalgae: a novelapproach for removing ammonium and phosphorus from municipalwastewater. Water Res. 38, 466–474.

Donmez, G., Aksu, Z., 2002. Removal of chromium(VI) from salinewastewaters by Dunaliella species. Process Biochem. 38, 751–762.

Donmez, G.C., Aksu, Z., Ozturk, A., Kutsal, T., 1999. A comparativestudy on heavy metal biosorption characteristics of some algae.Process Biochem. 34, 885–892.


Ertesvag, H., Valla, S., 1998. Biosynthesis and applications of alginates.Polym. Degrad. Stabil. 59, 85–91.

Faafeng, B.A., van Donk, E., Kallqvist, T., 1994. In situ measurement ofalgal growth potential in aquatic ecosystems by immobilized algae. J.Appl. Phycol. 6, 301–308.

Falkowsky, P.G., 1980. Primary productivity in the sea. In: Falkowski,P.G. (Ed.), Environmental Science Research, vol. 19. Plenum Press,New York/London.

Frense, D., Muller, A., Beckmann, D., 1998. Detection of environmentalpollutants using optical biosensor with immobilized algae cells. Sens.Accumul. B 51, 256–260.

Garbayo, I., Vigara, A.J., Conchon, V., Martins, R., Dos Santos, V.A.P.,Vilchez, C., 2000. Nitrate consumption alterations induced by alginate-entrapment of Chlamydomonas reindhartii cells. Process Biochem. 36,459–466.

Garbisu, C., Gil, J.M., Bazin, M.J., Hall, D.O., Serra, J.L., 1991. Removalof nitrate from water by foam-immobilized Phormidium laminosum inbatch and continuous-flow bioreactors. J. Appl. Phycol. 3, 221–234.

Garnham, G.W., Codd, G.A., Gadd, G.M., 1992a. Accumulation ofcobalt, zinc and manganese by the estuarine green microalga Chlorella

salina immobilized in alginate microbeads. Environ. Sci. Technol. 26,1764–1770.

Garnham, G.W., Codd, G.A., Gadd, G.M., 1992b. Kinetics of uptake andintracellular location of cobalt, manganese and zinc in the estuarinegreen alga Chlorella salina. Appl. Microbiol. Biotech. 37, 270–276.

Gee, A.R., Dudeney, A.W.L., 1987. Adsorption and crystallization ofgold at biological surfaces. In: International Symposium on Biohy-drometallurgy. Warwick, UK.

Ghosh, M., Gaur, J.P., 1998. Current velocity and the establishment ofstream algal periphyton communities. Aquat. Bot. 60, 1–10.

Greene, B., Bedell, G.W., 1990. Algal gels or immobilized algae for metalrecovery. In: Akatsuka, I. (Ed.), Introduction to Applied Phycology.SPB Academic Publishing BV, The Hague, The Netherlands, pp. 137–149.

Gualtieri, P., Barsanti, L., Passarelli, V., 1988. Chitosan as flocculant forconcentrating Euglena gracilis cultures. Ann. Inst. Pasteur Microbiol.139, 717–726.

Guo, P., Wang, J., Li, X., Zhu, J., Reinert, T., Heitmann, J., Spemann, D.,Vogt, J., Flagmeyer, R.H., Butz, T., 2000. Study of metal bioaccumu-lation by nuclear microprobe analysis of algae fossils and living algaecells. Nucl. Instrum. Methods B 161–163, 801–807.

Hashim, M.A., Tan, H.N., Chu, K.H., 2000. Immobilized marine algalbiomass for multiple cycles of copper adsorption and desorption. Sep.Purif. Technol. 19, 39–42.

Hatanaka, Y., Kudo, T., Miyataka, M., Kobayashi, O., Higashihara, M.,Hiyama, K., 1999. Asymmetric reduction of hydroxyacetone topropanediol in immobilized halotolerant microalga Dunaliella parva.J. Biosci. Bioeng. 88, 281–286.

Hertzberg, S., Jensen, A., 1989. Studies of alginate-immobilized marinemicroalgae. Bot. Mar. 32, 267–273.

Iqbal, M., Edyvean, R.G.J., 2004. Biosorption of lead, copper and zincions on loofa sponge immobilized biomass of Phanaerochaete chry-

sosporium. Miner. Eng. 17, 217–223.Iqbal, M., Edyvean, R.G.J., 2005. Loofa sponge immobilized fungal

biosorbent: a robust system for cadmium and other dissolved metalremoval from aqueous solution. Chemosphere 61, 510–518.

ISO, 1995. ISO 10253:1995(E). Water quality – Marine algal growthinhibition test with Skeletonema costatum and Phaeodactylum tricor-

nutum. pp. 1–8.Jang, L.K., 1993. Diffusivity of Cu2+ in calcium alginate gel beads.

Biotech. Bioeng. 43, 183–185.Jang, L.K., Nguyen, D.V., Kolostyak, K., Geesey, G.G., 1995a. Addition

of copper-sequestering agents to alginate gel to enhance copperrecovery from aqueous media. Water. Res. 29, 2525–2529.

Jang, L.K., Nguyen, D., Geesey, G.G., 1995b. Effect of pH on theabsorption of Cu(II) by alginate gel. Water. Res. 29 (1), 315–321.

Jang, L.K., Nguyen, D., Geesey, G.G., 1995c. Selectivity of alginate gelfor Cu vs. Co. Water. Res. 29 (1), 307–313.

Jen, A.C., Wake, M.C., Mikos, A.G., 1996. Review: hydrogels for cellimmobilization. Biotechnol. Bioeng. 50, 357–364.

Jeon, C., Park, J.Y., Yoo, Y.J., 2002. Novel immobilization of alginic acidfor heavy metal removal. Biochem. Eng. J. 11, 159–166.

Joo, D.S., Cho, M.G., Lee, J.S., Park, J.H., Kwak, J.K., Han, Y.H.,Bucholz, R., 2001. New strategy for the cultivation of microalgae usingmicroencapsulation. J. Microencapsul. 18, 567–576.

Jimenez-Perez, M.V., Sanchez-Castillo, P., Romera, O., Fernandez-Moreno, D., Perez-Martınez, C., 2004. Growth and nutrient removalin free and immobilized planktonic green algae isolated from pigmanure. Enzyme Microb. Tech. 34, 392–398.

Kadam, K.L., 2002. Environmental implications of power generation viacoal-microalgae cofiring. Energy 27, 905–922.

Kapdan, I.K., Kargi, F., 2006. Bio-hydrogen production from wastematerials. Enzyme Microb. Technol. 38, 569–582.

Kawabata, N., Nishimura, S., Yoshimura, T., 1990. New method ofimmobilization of microbial cells by capture on the surface of insolublePyridinium-type resin. Biotechnol. Bioeng. 35, 1000–1005.

Kaya, V.M., Picard, G., 1996. Stability of chitosan gel as entrapmentmatrix of viable Scenedesmus bicellularis cells immobilized onscreens for tertiary treatment of wastewater. Bioresour. Technol. 56,147–155.

Khattar, J.I.S., Sarma, T.A., Singh, D.P., 1999. Removal of chromiumions by agar immobilized cells of the cyanobacterium Anacystis

nidulans in a continuous flow bioreactor. Enzyme Microb. Technol. 25,564–568.

Khoshmanesh, A., Lawson, F., Prince, I.G., 1997. Cell surface area as amajor parameter in the uptake of cadmium by unicellular greenmicroalgae. Chem. Eng. J. 65, 13–19.

Kidambi, S.P., Sundin, G.W., Palmer, D.A., Chakrabarty, A.M., Bende,C.L., 1995. Copper as a signal for alginate synthesis in Pseudomonas

syringae pv. syringae. Appl. Environ. Microb. (June), 2172–2179.Kosakowska, A., Lewandowska, J., Ston, J., Burkiewicz, K., 2004.

Qualitative and quantitative composition of pigments in Phaeodacty-

lum tricornutum (Bacillariophyceae) stressed by iron. BioMetals 17,45–52.

Kubal, B.S., D’Souza, S.F., 2004. Immobilization of catalase by entrap-ment of permeabilized yeast cells in hen egg white using glutaralde-hyde. J. Biochem. Bioph. Methods 59, 61–64.

La Rosa, T., Mirto, S., Mazzola, A., Danovaro, R., 2001. Differentialresponses of benthic microbes and meiofauna to fish-farm disturbancein coastal sediments. Environ. Pollut. 112, 427–434.

Lau, P.S., Tam, N.F.Y., Wong, Y.S., 1998. Effect of carrageenanimmobilization on the physiological activities of Chlorella vulgaris.Bioresour. Technol. 63, 115–121.

Laurinavichene, T.V., Fedorov, A.S., Ghirardi, M.L., Seibert, M.,Tsygankov, A.A., 2006. Demonstration of sustained photoproductionby immobilized, sulphur deprived Chlamydomonas reindhartii cells. Int.J. Hydrogen Energy 31, 659–667.

Leon, R., Garbayo, I., Hernandez, R., Vigara, J., Vilchez, C., 2001.Organic solvent toxicity in photoautotrophic unicellular microorgan-isms. Enzyme Microb. Technol. 29, 173–180.

Liu, Y.K., Seki, M., Tanaka, H., Furusaki, S., 1998. Characteristics ofloofa (Luffa cylindrica) sponge as a carrier for plant cell immobiliza-tion. J. Ferment. Bioeng. 85, 416–421.

Lubian, L.M., 1989. Concentrating cultured marine microalgae withchitosan. Aquacult. Eng. 8, 257–265.

Lukavsky, J., 1988. Long-term preservation of algal strains by immobi-lization. Arch. Protistenkd. 135, 65–68.

Lukavsky, J., Komarek, J., Lukavska, A., Ludvık, J., Pokorny, J., 1986.Metabolic activity and cell structure of immobilized algal cells(Chlorella, Scenedesmus). Arch. Hydrobiol. Suppl. 73, 261–279.

Lukavsky, J., Marsalek, B., 1997. The evaluation of toxicity by abiosensor with immobilized algae. Algol. Stud. 85, 147–155.

Maeda, S., Sakaguchi, T., 1990. Accumulation and detoxification of toxicelements by algae. In: Akatsuka, I. (Ed.), Introduction to AppliedPhycology. SPB Academic Publishing BV, The Hague, TheNetherlands.


Malik, A., 2004. Metal bioremediation through growing cells. Environ.Int. 30, 261–278.

Mallick, N., 2002. Biotechnological potential of immobilized algaefor wastewater N, P and metal removal: a review. BioMetals 15,377–390.

Markov, S.A., Eivazova, E.R., Greenwood, J., 2006. Photostimulation ofH2 production in the green alga Chlamydomonas reindhartii uponphotoinhibition of its O2-evolving system. Int. J. Hydrogen. Energy 31,1314–1327.

Mayasich, J.M., Karlander, E.P., Terlizzi Jr., D.E., 1986. Growthresponses of Nannochloris oculata Droop and Phaeodactylum tricor-

nutum Bohlin to the herbicide atarazine as influenced by light intensityand temperature. Aquat. Toxicol. 8, 175–184.

Melis, A., 2002. Green alga hydrogen production: progress, challenges andprospects. Int. J. Hydrogen Energy 27, 1217–1228.

Moreira dos Santos, M., Moreno-Garrido, I., Goncalves, F., Soares,A.M.V.M., Ribeiro, R., 2002. An in situ bioassay with microalgae forestuarine environments. Environ. Toxicol. Chem. 21, 567–574.

Moreira dos Santos, M., Soares, A.M.V.M., Ribeiro, R., 2004. An in situbioassay for freshwater environments with the microalga Pseudokirch-

neriella subcapitata. Ecotox. Environ. Safe 59, 164–173.Moreira, S.M., Moreira-Santos, M., Guilhermino, L., Ribeiro, R., 2006.

Immobilization of the marine microalga Phaeodactylum tricornutum inalginate for in situ experiments: bead stability and suitability. EnzymeMicrob. Technol. 38, 135–141.

Morelli, E., Pratesi, E., 1997. Production of phytochelatins in the marinediatom Phaeodactylum tricornutum in response to copper and cad-mium exposure. Environ. Contam. Tox. B 59, 657–664.

Morelli, E., Scarano, G., 2001. Synthesis and stability of phytochelatinsinduced by cadmium and lead in the marine diatom Phaeodactylum

tricornutum. Mar. Environ. Res. 52, 383–395.Moreno-Garrido, I., Codd, G.A., Gadd, G.M., Lubian, L.M., 2002. Cu

and Zn accumulation by calcium alginate immobilized marine micro-algal cells of Nannochloropsis gaditana (EUSTIGMATOPHYCEAE).Cienc. Mar. 28, 107–119.

Moreno-Garrido, I., Campana, O., Lubian, L.M., Blasco, J., 2005.Calcium alginate immobilized marine microalgae: Experiments ongrowth and short-term heavy metal accumulation. Mar. Pollut. Bull.51, 823–929.

Moreno-Garrido, I., Blasco, J., Gonzalez-DelValle, M., Lubian, L.M.,1998. Differences in copper accumulation by the marine microalgaeNannochloropsis gaditana Lubian, submitted to two different thermaltreatments. Ecotox. Environ. Restor. 1, 43–47.

Moreno-Garrido, I., Lubian, L., Blasco, J., 2007. Sediment toxicity testsinvolving immobilized microalgae (Phaeodactylum tricornutum Boh-lin). Environ. Int. 33, 481–495.

Munawar, M., Munawar, I.F., 1987. Phytoplankton bioassays forevaluating toxicity of in situ sediment contaminants. Hydrobiologia149, 87–105.

Munoz, R., Alvarez, M.T., Munoz, A., Terrazas, E., Guieysse, B.,Mattiasson, B., 2006. Sequential removal of heavy metals ions andorganic pollutants using an algal–bacterial consortium. Chemosphere63, 903–911.

Munoz, R., Guieysse, B., 2006. Algal–bacterial processes for the treatmentof hazardous contaminants: a review. Water Res. 40, 2799–2815.

Naessens, M., Leclerc, J.C., Tran-Minh, C., 2000. Fiber optic sensor usingChlorella vulgaris for determination of toxic compounds. Ecotox.Environ. Safe 46, 181–185.

Nagase, H., Pattanasupong, A., Sugimoto, E., Tani, K., Nasu, M., Hirata,K., Miyamoto, K., 2006. Effect of environmental factors on perfor-mance of immobilized consortium system for degradation of carbeda-zim and 2,4-dichlorophenoxyacetic acid in continuous culture.Biochem. Eng. J. 29, 163–168.

Nakasaki, K., Murai, T., Akiyama, T., 1989. Dynamic modelling of animmobilized cell reactor. Appl. Biochem. Biotech. 22, 279–289.

Nayar, S., Goh, B.P.L., Chou, L.M., 2005. Settlement of marineperiphytic algae in a tropical estuary. Estuar. Coast. Shelf S. 64,241–248.

Nayar, S., Goh, B.P.L., Chou, L.M., Reddy, S., 2003. In situ microcosmsto study the impact of heavy metals resuspended by dredging onperiphyton in a tropical estuary. Aquat. Toxicol. 64, 293–306.

Nestle, N., Kimmich, R., 1996. NMR microscopy of heavy metalabsorption in calcium alginate beads. Appl. Biochem. Biotechnol. 56,9–17.

Nowak, E.C.M., Podola, B., Melkonian, M., 2005. The 96-well twin-layersystem: a novel approach in the cultivation of microalgae. Protist 156,239–251.

Ogbonn, J.C., Tomiyama, S., Tanaka, H., 1996. Development of a methodfor immobilization of non-flocculating cells in loofa (Luffa cylindrica)sponge. Process Biochem. 31, 737–744.

Oh, Y.S., Maeng, J., Kim, S.J., 2000. Use of microorganism-immobilizedpolyurethane foams to adsorb and degrade oil on water surface. Appl.Microb. Biotechnol. 54, 418–423.

Olguın, E.J., 2003. Phycoremediation: key issues for cost-effective nutrientremoval processes. Biotechnol. Adv. 22, 81–91.

Oungbho, K., Muller, B.W., 1997. Chitosan sponges as sustain releasedrug carriers. Int. J. Pharm. 156, 229–237.

Overnell, J., 1975. The effect of heavy metals on photosynthesis and loss ofcell potassium in two species of marine algae, Dunaliella tertiolecta andPhaeodactylum tricornutum. Mar. Biol. 29, 99–103.

Pane, L., Feletti, M., Bertino, C., Carli, A., 1998. Viability of the marinemicroalga Tetraselmis suecica grown free and immobilized in alginatebeads. Aquacult. Int. 6, 411–420.

Papageorgiou, G.C., 1987. Immobilized photosynthetic microorganisms.Photosynthetica 21, 367–383.

Pavlic, Z., Vidakovic-Cifrek, Z., Puntaric, D., 2005. Toxicity of surfac-tants to green microalgae Pseudokirchneriella subcapitata and Scene-

desmus subspicatus and to marine diatoms Phaeodactylum tricornutum

and Skeletonema costatum. Chemosphere 61, 1061–1068.Podola, B., Nowack, E.C.M., Melkonian, M., 2004. The use of multiple-

strain algal sensor chips for the detection and identification of volatileorganic compounds. Biosens. Bioelectron. 19, 1253–1260.

Polle, J.E.W., Kanakagiri, S., Jin, E.S., Masuda, T., Melis, A., 2002.Truncated chlorophyll antenna size of the photosystems – a practicalmethod to improve microalgal productivity and hydrogen productionin mass culture. Int. J. Hydrogen Energy 27, 1257–1264.

Poncelet, D., Babak, V., Dulieu, C., Picot, A., 1999. A physico-chemicalapproach to production of alginate beads by emulsification-internalionotropic gelation. Coll. Surf. A: Physicochem. Eng. Asp. 155, 171–176.

Prieto, M.B., Hidalgo, A., Serra, J.L., Llama, M.J., 2002. Degradation ofphenol by Rhodococcus erythropolis UPV-1 immobilized on Biolite� ina packed-bed reactor. J. Biotechnol. 97, 1–11.

Proulx, D., de la Noue, J., 1988. Removal of macronutrients fromwastewaters by immobilized microalgae. In: Moo-Young, M. (Ed.),Bioreactor Immobilized Enzymes and Cells. Fundamentals andApplications. Elsevier Applied Science Publ. Ltd., Essex.

Quong, D., Neufeld, R.J., Skjak-Bræk, G., Poncelet, D., 1998. Externalversus internal source of calcium during the gelation of alginate beadsfor DNA encapsulation. Biotechnol. Bioeng. 57, 438–446.

Radix, P., Leonard, M., Papantoniou, C., Roman, G., Saouter, E.,Gallotti-Schmitt, S., Thiebaud, H., Vasseur, P., 2000. Comparison offour chronic toxicity tests using algae, bacteria and invertebrates withsixteen chemicals. Ecotox. Environ. Safe 47, 186–194.

Radwan, S.S., Al-Hasan, R.H., Salamah, S., Al-Dabbous, S., 2002.Bioremediation of oily sea water by bacteria immobilized in biofilmscoating macroalgae. Int. Biodeter. Biodegr. 50, 55–59.

Rangasayatorn, N., Pokethitiyook, P., Upatahm, E.S., Lanza, G.R., 2004.Cadmium biosorption by cells of Spirulina platensis TISTR 8217immobilized in alginate and silica gel. Environ. Int. 30, 57–63.

Robinson, P.K., Mak, A.L., Trevan, M.D., 1986. Immobilized algae: areview. Process Biochem. (August), 122–126.

Robinson, P.K., Reeve, J.O., Goulding, K.H., 1988. Kinetics of phos-phorus uptake by immobilized Chlorella. Biotechnol. Lett. 10, 17–20.

Romo, S., Perez-Martınez, C., 1997. The use of immobilization in alginatebeads for long-term storage of Pseudanabaena galeata (Cyanobacteria)in the laboratory. J. Phycol. 33, 1073–1076.


Sanders, C.A., Rodrıguez Jr., M., Greenbaun, E., 2001. Stand-off tissue-based biosensors for the detection of chemical warfare agents usingphotosynthetic fluorescence induction. Biosens. Bioelectron. 16, 439–446.

Santos-Rosa, F., Galvan, F., Vega, J.M., 1989. Photoproduction ofammonium by Chlamydomonas reinhardtii cells immobilized in bariumalginate: a reactor feasibility study. Appl. Microbiol. Biotechnol. 32,285–290.

Schreiter, P.P.Y., Gillor, O., Post, A., Belkin, S., Schmid, R.D.,Bachmann, T.T., 2001. Monitoring of phosphorus bioavailability inwater by an immobilized luminescent cyanobacterial reporter strain.Biosens. Bioelectron. 16, 811–818.

Seki, H., Suzuki, A., 2002. Adsorption of heavy metal ions to floc-typebiosorbents. J. Coll. Interf. Sci. 249, 295–300.

Semple, K.T., 1998. Heterotrophic growth on phenolic mixtures byOchromonas danica. Res. Microbiol. 149, 65–72.

Semple, K.T., Cain, R.B., 1996. Biodegradation of phenols by the algaOchromonas danica. Appl. Environ. Microb. 62, 1265–1273.

Shitanda, I., Takada, K., Sakai, Y., Tatsuma, T., 2005. Compactamperometric algal biosensors for the evaluation of water toxicity.Anal. Chim. Acta 530, 191–197.

Singh, R., Prasad, B.B., 2000. Trace metal analysis: selective sample(copper II) enrichment on an AlgaSORB column. Process Biochem.35, 897–905.

Smidsrød, O., Skjak-Braek, G., 1990. Alginate as immobilization matrixfor cells. Tibtech 8, 71–78.

Spinti, M., Zhuang, H., Trujillo, E.M., 1995. Evaluation of immobilizedbiomass beads for removing heavy metals from wastewaters. WaterEnviron. Res. 67, 943–952.

Stark, P.C., Rayson, G.D., 2000. Comparisons of metal–ion binding toimmobilized biogenic materials in a flowing system. Adv. Environ.Res. 4, 113–122.

Streble, H., Krauter, D., 1987. Atlas de los microorganisms de agua dulce(Atlas of Freshwater Microorganisms). Omega S.A., Barcelona.

Suzuki, T., Yamaguchi, T., Ishida, M., 1998. Immobilization of Proto-

theca zopfii in calcium alginate vedas for the degradation of hydro-carbons. Process Biochem. 33, 541–546.

Tam, N.F.Y., Wong, Y.S., 2000. Effect of immobilized microalgal beadconcentrations on wastewater nutrient removal. Environ. Pollut. 107,145–151.

Thakur, A., Kumar, H.D., 1999. Nitrate, ammonium and phosphateuptake by the immobilized cells of Dunaliella salina. Environ. Contam.Tox. B 62, 70–78.

Thepenier, C., Gudin, C., Thomas, D., 1985. Immobilization of Porphyri-

dium cruentum in polyurethane foams for the production of polysac-charide. Biomass 7, 225–240.

Tosa, T., Sato, T., Mori, T., Yamamoto, K., Takata, I., Nishida, Y.,Chibata, I., 1979. Immobilization of enzymes and microbial cells usingcarrageenan as matrix. Biotechnol. Bioeng. 21, 1697–1709.

Travieso, L., Pellon, A., Benıtez, F., Sanchez, E., Borja, R., O’Farrill,N.O., Weiland, P., 2002. BIOALGA reactor: preliminary studies forheavy metals removal. Biochem. Eng. J. 12, 87–91.

Travieso, L., Benıtez, F., Weiland, P., Sanchez, E., Dupeyron, R.,Domınguez, A.R., 1996. Experiments on immobilization of microalgaefor nutrient removal in wastewater treatments. Bioresour. Technol. 55,181–186.

Travieso Cordoba, L., Sanchez Hernandez, E., 1995a. Final treatment forcattle manure using immobilized microalgae. II. Influence of therecirculation. Resour. Conserv. Recy. 13, 177–182.

Travieso Cordoba, L., Sanchez Hernandez, E., Weiland, P., 1995b. Finaltreatment for cattle manure using immobilized microalgae. I. Study ofthe support media. Resour. Conserv. Recy. 13, 167–175.

Tripathi, U., Ramachandra, R.S., Ravishankar, G.A., 2002. Biotrans-formation of phenylpropanoid compounds to vanilla flavormetabolites in cultures of Haematococcus pluvialis. Process Biochem.38, 419–426.

Twist, H., Edwards, A.C., Codd, G.A., 1997. A novel In situ biomonitorusing alginate immobilised algae (Scenedesmus subspicatus) for theassessment of eutrophication in flowing surface waters. Water Res. 31,2066–2067.

Urrutia, I., Serra, J.L., Llama, M.J., 1995. Nitrate removal from water byScenedesmus obliquus immobilizes in polymeric foams. Enzyme Mic-rob. Technol. 17, 200–205.

Valdman, E., Erijman, L., Pessoa, F.L.P., Leite, S.G.F., 2001. Continuousbiosorption of Cu and Zn by immobilized waste biomass Sargassum

sp. Process Biochem. 36, 869–873.van Vuuren, D.P., Cofala, J., Eerens, H.E., Oostenrijk, R., Heyes, C.,

Klimont, Z., den Elzen, M.G.J., Amann, M., 2006. Exploring theancillary benefits of the Kyoto Protocol for air pollution in Europe.Energ. Policy 34, 444–460.

Vignoli, J.A., Celligoi, M.A.P.C., Silva, R.S.F., 2006. Development of astatistical model for sorbitol production by free and immobilizedZymomonas mobilis in loofa sponge Luffa cylindrica. Process Biochem.41, 240–243.

Vılchez, C., Vega, J.M., 1994. Nitrite uptake by Chlamydomonas

reinhardtii cells immobilized in calcium alginate. Appl. Microbiol.Biotechnol. 41, 137–141.

Vılchez, C., Vega, J.M., 1995. Nitrite uptake by immobilized Chlamydo-

monas reinhardtii cells growing in airlift reactors. Enzyme Microb.Technol. 17, 386–390.

Vılchez, M.J., Vigara, J., Garbayo, I., Vılchez, C., 1997. Electronmicroscopic studies on immobilized growing Chlamydomonas rein-

hardtii cells. Enzyme Microb. Technol. 21, 25–47.Vılchez, C., Garbayo, I., Markvicheva, E., Galvan, F., Leon, R., 2001.

Studies on the suitability of alginate-entrapped Chlamydomonas

reindhartii cells for sustaining nitrate consumption processes. Biore-sour. Technol. 78, 55–61.

Weller, M.G., 2000. Immunochromatographic techniques – a criticalreview. Fresen. J. Anal. Chem. 366, 635–645.

Widerøe, H., Danielsen, S., 2001. Evaluation of the use of Sr2+ in alginateimmobilized cells. Naturwissenchaften 88, 224–228.

Wiegman, S., Termeer, J.A.G., Verheul, T., Kraak, M.H.S., De Voogt, P.,Laane, R.W.P.M., Admiraal, W., 2002. UV absorbance dependenttoxicity of acridine to the marine diatom Phaeodactylum tricornutum.Environ. Sci. Technol. 36, 908–913.

Wijffels, R.H., Tramper, J., 1989. Performance of growing Nitrosomonas

europea cells immobilized in j-carrageenan. Appl. Microbiol. Biotech-nol. 32, 108–112.

Willke, B., Willke, T., Vorlop, K.D., 1994. Poly(carbamoylsulphonate) asmatrix for whole cell immobilization – biological characterization.Biotechnol. Tech. 8, 623–626.

Yamaguchi, T., Ishida, M., Suzuki, T., 1999. An immobilized cell systemin polyurethane foam for the lipophilic micro-alga Prototheca zopfii.Process Biochem. 34, 167–171.

Zeglinska, L., 2005. Personal communication. Institute of Oceanology,Polish Academy of Sciences, Powstancow Warszawy 55 (81-712 Sopot,Poland).

spi

Documents