ARTICLE

The Microalga Parachlorella kessleri––A NovelHighly Efficient Lipid Producer

Xiuling Li,1 Pavel Pribyl,2 Katerina Bisova,1 Shigeyuki Kawano,3 Vladislav Cepak,2

Vilem Zachleder,1 Maria Cızkova,1 Irena Branyikova,1 Milada Vıtova1

1Laboratory of Cell Cycle of Algae, Institute of Microbiology, AS CR, Opatovicky mlyn,

379 81 Trebon, Czech Republic; telephone: þ420-384-340-480; fax: þ420-384-340-415;

e-mail: zachleder@gmail.com2Institute of Botany, AS CR, Algological Centre, Trebon, Czech Republic3Department of Integrated Sciences, Graduate School of Frontier Sciences,

University of Tokyo, Kashiwanoha, Kashiwa, Chiba, Japan

ABSTRACT: The alga Parachlorella kessleri, strain CCALA255, grown under optimal conditions, is characterized bystorage of energy in the form of starch rather than lipids. Ifgrown in the complete medium, the cultures grew rapidly,producing large amounts of biomass in a relatively shorttime. The cells, however, contained negligible lipid reserves(1–10% of DW). Treatments inducing hyperproduction ofstorage lipids in P. kessleri biomass were described. Thecultures were grown in the absence or fivefold decreasedconcentration of either nitrogen or phosphorus or sulfur.Limitation by all elements using fivefold or 10-fold dilutedmineral medium was also tested. Limitation with anymacroelement (nitrogen, sulfur, or phosphorus) led to anincrease in the amount of lipids; nitrogen limitation was themost effective. Diluted nutrient media (5- or 10-fold) wereidentified as the best method to stimulate lipid overproduc-tion (60% of DW). The strategy for lipid overproductionconsists of the fast growth of P. kessleri culture grown in thecomplete medium to produce sufficient biomass (DWmorethan 10 g/L) followed by the dilution of nutrient medium tostop growth and cell division by limitation of all elements,leading to induction of lipid production and accumulationup to 60% DW. Cultivation conditions necessary for maxi-mizing lipid content in P. kessleri biomass generated in ascale-up solar open thin-layer photobioreactor weredescribed.

Biotechnol. Bioeng. 2013;110: 97–107.

� 2012 Wiley Periodicals, Inc.

KEYWORDS: carbon dioxide; light intensity; limitation byelements; lipid hyperproduction; Parachlorella kessleri;thin-layer photobioreactor

Introduction

Considering natural resource depletion and climate change,lipid-based algal biofuels are an alternative to conventionalfossil fuels, due to the high productivity of algal biomass(Chisti, 2007; Doucha and Lıvansky, 2009; Rodolfi et al.,2009; Singh et al., 2011b) and the ability of algae torecycle CO2 originating from flue gas (Benemann, 1997;Brown, 1996; Douskova et al., 2009). Algae are considered asthe only alternative to current biofuel crops such as corn andsoybean, as they do not require arable land (Chisti, 2007; Huet al., 2008; Singh et al., 2011a).

Increased interest has been focused on microalgae not onlydue to their high growth rate and high photosyntheticefficiency, but particularly due to the possibility of controllingtheir metabolism to produce relatively high contents ofenergy-rich compounds, either starch (Branyikova et al.,2011) and/or lipids (Chen et al., 2011; Deng et al., 2009; Lee,2011). Under favorable growth conditions, algae synthesizefatty acids, principally as substrates for esterification intoglycerol-based polar lipids (e.g. glycolipids and phospholi-pids), the main constituent of membranes. However, underunfavorable environmental or stress conditions, many algaealter their lipid biosynthetic pathways towards the synthesisand accumulation of neutral lipids (20–50% cell dry weight),mainly in the form of triacylglycerols (Hu et al., 2008), thatare typically stored as cytoplasmic lipid bodies (Goodsonet al., 2011; Yu et al., 2007). Neutral lipids can be readilyconverted to biodiesel or other fuel types through both

Correspondence to: V. Zachleder

Contract grant sponsor: Ministry of Education, Youth and Sports of the Czech Republic

Contract grant number: OE09025; LH12145

Contract grant sponsor: Grant Agency of the Czech Republic

Contract grant number: P503/10/1270; P501/10/P258

Contract grant sponsor: CREST of Japan Science and Technology Agency

Contract grant sponsor: Technology Agency of the Czech Republic

Contract grant number: TE 01020080

Contract grant sponsor: Academy of Sciences of the Czech Republic

Contract grant number: RVO 61388971

Received 2 April 2012; Revision received 4 june 2012, Accepted 22 june 2012

Accepted manuscript online 5 July 2012;

Article first published online 18 July 2012 in Wiley Online Library

(http://onlinelibrary.wiley.com/doi/10.1002/bit.24595/abstract)

DOI 10.1002/bit.24595

� 2012 Wiley Periodicals, Inc. Biotechnology and Bioengineering, Vol. 110, No. 1, January, 2013 97

existing and emerging lipid refining processes (Biller andRoss, 2011; Francisco et al., 2010; Halim et al., 2011; Patilet al., 2011). However, oleaginous microalgal strains thathave been used to date often showedmuch lower growth ratesthan many non-oleaginous species (Hu et al., 2008; Sheehanet al., 1998), leading to unacceptably expensive biofuelproduction.

Previous studies have shown that some microalgae (e.g.selected strains of Chlorella) can be directed to accumulatehigh levels of starch as a storage compound; this may be usedfor industrial production of bioethanol (Branyikova et al.,2011). Several approaches like inhibitory treatments bycycloheximide or fluorodeoxyuridine, or limitation ofmacroelements, such as phosphorus (Ballin et al., 1988),sulfur (Setlık et al., 1988), or nitrogen (Zachleder et al.,1988), were used to stimulate overproduction of starch inmicroalgae (Branyikova et al., 2011).

Although the mechanisms of induction of lipid produc-tion can be different from starch, there are several commonapproaches inducing both starch and lipid overproduction.It is widely believed that the lipid content could be increasedby nitrogen or phosphate limitation (Mutlu et al., 2011;Reitan et al., 1994; Tornabene et al., 1983). Currently,nitrogen limitation is the most frequently used treatment toenhance lipid production in microalgae (Rodolfi et al.,2009). It was also shown that the algal strains appropriate foroverproduction of starch are not usually suitable foroverproduction of lipids and vice versa (Branyikova et al.,2011; Li et al., 2010a,c).

Many algae, particularly green algae (Chlorophyta), usestarch as the primary carbon and energy storage compoundand in Chlamydomonas reinhardtii under nitrogen deficien-cy (�N) conditions, its content can reach up to 45% of celldry weight (Li et al., 2010a) or higher (60% of DW) in someChlorella strains if induced by sulfur deficiency or inhibitedby cycloheximide (Branyikova et al., 2011). Some algae mayaccumulate similar amounts of starch and lipids under stressconditions (Li et al., 2011; Ramazanov and Ramazanov,2006), whereas others only transiently accumulate starch(Collen et al., 2004).

Substantial differences in both biomass and lipidproductivities as well as in final content of neutrallipids can be found among various Chlorella strains.Recently, a highly productive strain, Chlorella vulgarisCCALA 256 (Pribyl et al., 2012) was identified anddescribed. Another promising strain, Parachlorella kessleriCCALA 255, characterized by high biomass and lipidproductivity (Pribyl et al., 2012), is being presented in detailin this study.

In the present study, the controlled production of oilreserves was tested under conditions of the absence orlimitation of some macroelements (nitrogen, sulfur, andphosphorus) as well as limitation of all elements in mineralmedia by dilution. To confirm the laboratory findingsfor potential industrial use, lipid production was testedin algal cultures grown in an outdoor scale-up thin layerphotobioreactor.

Materials and Methods

Microorganism and Culturing

The green microalga P. kessleri (Krienitz et al., 2004),strain CCALA 255, was provided by the Culture Collectionof Autotrophic Organisms (CCALA) in Trebon, CzechRepublic (http://www.butbn.cas.cz/ccala/index.php). In thecollection, the strain has been maintained on agar slantsunder irradiance of about 23mmol/(m2 s), 12/12 h (light/dark) regime and at a temperature of 12–158C.

Laboratory cultivation units consisted of glass cylinders(inner diameter 36mm, height 500mm), which were placedin a thermostatic bath (308C) and continuously illuminatedby a panel dimmable fluorescent lamps (OSRAM DULUXL55W/950 Daylight, Italy) allowing adjustment of theincident light intensity from 16 to 780mmol/(m2 s). Thecylinders were ‘‘aerated’’ using a mixture of air and CO2

(2%, v/v). The volume of the algal suspension in eachcylinder was 300mL, and each cylinder was supplied withgas at a flow rate of 15 L/h.

For the mineral medium used for experimental cultures,and a description of both laboratory and scale up outdoorcultivation units see Branyikova et al. (2011).

Nutrient Limitation

P. kessleri cultures were grown in complete medium to attainthe required cell concentration for any given experiment(usually between 0.3 and 0.75 g of DW/L). The originalmineral medium was removed from the cell suspension bycentrifugation and the cells were re-suspended in freshmineral medium (control) or in one of the nutrient-limitedmedia (for more details, see Branyikova et al., 2011).

Biomass Determination

For dry weight determination, biomass was separated fromthe medium by centrifugation of 2mL of the cell suspensionin pre-weighed microtubes at 3,000g for 5min; the sedimentwas dried at 1058C for 12 h and weighed on an analyticalSartorius 1601 MPB balance (as described in Branyikovaet al., 2011).

Cell volume and concentration were measured using aBeckman Coulter Multisizer III (Coulter Corporation,Miami, FL) by diluting 10–50mL of fixed (0.2% glutaralde-hyde) cell suspension into 10mL of 0.9% NaCl (w/v)electrolyte solution.

Measurement of Light Intensity

A quantum/radiometer-photometer (LI-COR, Inc.) wasused. In the culture unit, dimmable fluorescent tubes wereused for adjustment of irradiance. The mean light intensity(I) was calculated according to the Lambert–Beer equation:I¼ (Ii� It)/ln(Ii/It), where Ii is the incident light intensity

98 Biotechnology and Bioengineering, Vol. 110, No. 1, January, 2013

measured at the surface of the culture vessel and It is thetransmitted light intensity measured at the rear side ofculture vessel.

Determination of Chlorophyll Content

The algal suspension (10mL) was centrifuged at 4,000g for3min and the sediment was collected. Phosphate buffer,7.7 pH (1mL), a pinch of MgCO3, and Zircon beads(500mL, size 0.7) were added to the sediment, which wasthen disintegrated by vortexing (Vortex Genie 2, ScientificIndustries, Inc., Bohemia, NY) for 10min. Acetone (4mL,100%) was added, mixed well, and centrifuged at 4,000g for3min. The supernatant was drained into a calibrated testtube using an exhauster/air pump, closed with a stopper andleft standing in a dark-block. Another 4mL of acetone(80%) were added to the sediment, mixed well, andcentrifuged at 4,000g for 3min. Using an exhauster/airpump, the supernatant was drained off to the samecalibrated test tube used in the preceding step and toppedup with 80% acetone up to 10mL. Absorbances at 750, 664,647, 470, 450 nm were measured in a 1 cm path lengthcuvette using the spectrophotometer (Shimadzu UV-1800S). Calculation of chlorophyll content was based onabsorbances at different wavelengths and was carried outaccording to equations published previously (MacKinney,1941).

Starch Analysis

Amodification of the method of (McCready et al., 1950) wasused as described previously (Branyikova et al., 2011).

Visualization of Lipids Using Nile Red Fluorescence

Intracellular lipid droplets were stained using the neutral lipid-specific dye, Nile Red (9-diethylamino-5H-benzo(a)phenox-azine-5-one), following the protocol described earlier(Eltgroth et al., 2005) with slight modifications. Briefly,1mL of the cell suspension was fixed with glutaraldehyde at afinal concentration of 0.25% (v/v) and stained with 4mL ofNile Red (Sigma, N3013) stock solution (0.5mg/mL ofacetone) that was stored in the dark at 48C.

Samples were observed after 5min using an epifluorescenceOLYMPUS BX 51 microscope equipped with the filtercombination U-MNU2 (360–370nm excitation and>420nmemission). Photomicrographs were taken with a digital cameraDP72, and processed using Adobe Photoshop 7.

Lipid Analyses

Gravimetric Lipid Determination

Analyses have been carried out as described previously(Pribyl et al., 2012).

Nile Red Fluorescence Determination of Lipids

The algal suspension was fixed with glutaraldehyde to a finalconcentration of 0.25% (v/v) and loaded into wells (100mLper well) of a 96-well plate. Four microlitre of Nile Redsolution (were added to wells, fluorescence intensity wasmeasured using a 96-well plate luminometer (Tecan infinite200, Switzerland) with the following filters: excitation485 nm (bandwidth 20 nm), emission 595 nm (bandwidth10 nm).

Glyceryl trioleate (Sigma, T7140) was used as a lipidstandard to obtain a calibration curve. Fluorescenceintensity was compared to the total lipid content determinedgravimetrically to quantify the relative fluorescence intensityvalues.

Fatty Acids in Lipids

This analysis was carried out in the Laboratory of FungalGenetics and Metabolism, Institute of Microbiology, ASCRin Prague as described by Rezanka et al. (2010).

Results

Effect of Nutrient Limitation

When cultures of P. kessleri were grown in the completemedium under optimal growth conditions (308C, 2% CO2,incident light intensity of 780mmol/(m2 s)), they rapidlygrew, and their biomass reached approx. 8.0 g of DW/L in4 days (Fig. 1). However, accumulation of lipids was veryslow. No lipid bodies were found in cells until at least 2 daysof culture, and only small lipid bodies were observed in somecells of 5-day culture (Fig. 2). The relative content of lipids inthe cells of 4-day culture was not more than 10% of DW(Fig. 3D). Starch content reached a maximum of approx.15% in 1-day, thereafter gradually decreased to 10% during4 days of culture period (Fig. 3D).

When nitrogen, sulfur, or phosphorus was eliminatedfrom the complete medium, the growth rate severelydecreased, especially in the case of nitrogen free medium(Fig. 1A). Their biomass of 4-day culture was<2.0 g of DW/L. In contrast, the elimination of any of the aboveelements caused an accumulation of lipids. Elimination ofnitrogen induced a very fast lipid accumulation (Fig. 2).Small lipid bodies were observed in the cells of 1-day culture,and lipid body number increased in the course of culture.The lipid bodies fused together, and the most of theintracellular space was occupied by large lipid bodies in 5days. The elimination of phosphorus also induced a lipidaccumulation, and the cells of 5-day culture accumulatedlipid at nearly the same extent as under nitrogenelimination. Although the elimination of sulfur induced alipid accumulation, the quantity of accumulated lipids wasless than under elimination of nitrogen or phosphorus.Sulfur elimination had another undesirable effect. The

Li et al.: Alga Parachlorella kessleri for Lipid Production 99

Biotechnology and Bioengineering

number of dead cells (blue cells in Fig. 2) rapidly increasedwith cultivation time.

For the avoidance of the undesirable effects of lack of theabove elements, nitrogen, sulfur, or phosphorus wasdecreased to 20% of the complete medium instead ofthe elimination of them. The cultures slowly grew, andthe biomass in 4 days reached 2.4–3 g of DW/L (Fig. 1). Thedecrease in nitrogen caused an accumulation of lipids, andthe relative content of lipids reached 45% in 4 days (Fig. 3E).On the other hand, changes in starch content were noteffected by the decrease in nitrogen except for the slightlowering of the content. An incubation in a mediumdiluted to 20% or 10% of the complete medium was alsoreduced the growth (Fig. 1). The most effective accumula-tion of lipids was achieved by the cultures grown inthe medium diluted to 10%, with the relative content of55% (Fig. 3F).

Effect of CO2 Limitation

The optimal growth of the cultures required aeration with2% CO2. The limitation of CO2 by aeration with air causedthe decrease in growth (Fig. 3). Decrease in growth by CO2

limitation occurred also under a condition of nitrogenlimitation to 20% of the complete medium. However, nodifference in growth was observed between 2% CO2 and airwhen the cultures were grown in the medium diluted to 10%of the complete medium. No significant differences in thetime course of starch content were observed betweencultures grown with 2% CO2 and air, regardless of nutrientconditions. In contrast, the rates of the accumulation oflipids were decreased by CO2 limitation under conditions ofnutrient limitation. Fluorescence microscopic observationalso demonstrated the decrease in lipid accumulationby CO2 limitation (Fig. 4). The intracellular space of thecells grown with air-aeration was less occupied by lipidbodies than that of the cells grown with 2% CO2-aeration.

The total chlorophyll content was also affected by CO2

limitation. As for the cultures grown in the completemedium, the total chlorophyll content reached a maximumin 3 days and slowly decreased thereafter under thecondition of sufficient CO2 supply (Fig. 5). However,under the condition of CO2 limitation, the total chlorophyllcontent reached a maximum in 1-day, and soon decreasedeven though the cultures still continued to increase inbiomass (Fig. 3A). As for the cultures grown underconditions of nutrient limitation, the total chlorophyllcontent decreased from the start of culture, and the decreaserates were made faster by CO2 limitation.

Effect of Mean Irradiance on Lipid Yield

The cultures started to be grown with the mean irradiancesat 125, 160, 183, 216, and 292mmol/(m2 s) by theadjustment of the initial biomass at 4.0, 2.0, 1.0, 0.5, and0.25 g DW/L. The cultures of an initial biomass below 2.0 g/Lstarted to accumulate lipids 2 days after inoculation, andcontinued for 2–3 days (Fig. 6). The relative lipid contentreached approx. 50%. However, the cultures of an initialbiomass of 2.0 and 4.0 g/L delayed to start accumulation oflipids for 1-day, and their relative lipid contents and lipidproductivities were substantially lower (Table I), indicatingthat the mean irradiance was insufficient to lipid produc-tion. The sufficient production of lipids was carried out withthe mean irradiances beyond 125mmol/(m2 s) at which thecultures of an initial biomass of 1.0 g/L reached themaximum of relative lipid content.

Fatty Acid Composition

The fatty acid composition of the cultures grown underdifferent conditions was analyzed. Although the lipids weremainly composed of fatty acids of C16 and C18 under allconditions tested, the ratio of saturated and unsaturated

Figure 1. Variation in biomass concentration (DWg/L) in cultures of Parachlor-

ella kessleri. Cultures were grown in complete mineral medium (1 medium) (A and B),

or in various element limiting growth media as follows: nitrogen (�N), phosphorus

(�P), sulfur (�S) free media, or 10-fold diluted complete mineral medium (0.1 medium)

(A), fivefold lower content of either nitrogen (0.2 N), phosphorus (0.2 P), or sulfur (0.2 S)

media, or fivefold diluted complete mineral medium (0.2 medium) (B). The cultures

were cultivated in a laboratory photobioreactor under continuous illumination of an

incident light intensity of 780mmol/(m2 s) and at the same initial biomass concentra-

tions (0.3 g/L).

100 Biotechnology and Bioengineering, Vol. 110, No. 1, January, 2013

fatty acids altered by culture conditions (Table II). Whencultures were grown in the complete medium under theoptimal growth condition, the ratio of saturated, andunsaturated fatty acids was 0.917. The lipids of the culturesgrown under nutrient limitation were rich in unsaturatedfatty acids. The ratio of saturated and unsaturated fatty acidsdecreased to 0.55 under the best condition for lipidaccumulation, that is, 10-fold diluted medium.

Large Scale Industrial Thin-Layer Bioreactor

The cultures were incubated in the complete medium put ina large-scale thin-layer bioreactor to simulate the industrialproduction of lipid-rich biomass. Unfortunately climaticconditions during cultivation were unfavorable, low light

intensities, and temperatures, so that the cultures grew atabout half the rate of the optimal growth conditions in thelaboratory (Fig. 7). Starch content decreased to 2% in thefirst 4 days, and thereafter increased gradually. It tookanother 8 days to reach approx. 15%, corresponding to thevalues in the laboratory.

On the 14th day from the beginning of the experiment,the 10-fold diluted medium was added to the reactor. Anaccumulation of lipids occurred 4 days after the addition,and the lipid content reached a maximum in next 3 days. Itwas slightly delayed compared to laboratory conditions,probably resulting from insufficient light intensity. Themeasurements of mean light intensity showed that thecultures in the thin-layer bioreactor efficiently absorbedlight, resulting in that the biomass in the outdoor thin-layerbioreactor reached almost 2 times higher (14 g/L) than in the

Figure 2. Time course of lipids accumulation during batch cultivation of P. kessleri. Lipid bodies were stained using Nile Red (yellow); autofluorescence of chloroplasts is

seen in red. Cultures were grown in complete mineral medium (1 medium), or in either nitrogen (�N), phosphorus (�P) or sulfur (�S) free media. The cultivation time for all variants

is indicated in the left column. The same initial biomass concentration (0.3 g/L) was used for all cultures and no detectable lipid bodies were seen in cells at the beginning (0 h; not

shown in figure). Scale bar¼ 10mm.

Li et al.: Alga Parachlorella kessleri for Lipid Production 101

Biotechnology and Bioengineering

laboratory bioreactor (8 g/L) even though the incident lightintensity of outdoor conditions was lower. The lipid contentof the outdoor cultures reached 25%, comparable to thelaboratory cultures of an initial biomass of 2.0 g/L (Fig. 6).

Discussion

Algae producing starch as a primary energy storagecompound usually have a relatively low lipid content.The starch reserves can be increased markedly by macro-element limitation in some algal species or strains such asScenedesmus quadricauda (Ballin et al., 1988; Setlık et al.,1988; Zachleder et al., 1988), C. vulgaris strain CCALA 924(Branyikova et al., 2011) or Tetraselmis subcordiformis (Yao

et al., 2012). A high starch content (about 40–45% of DW)was also found under nitrogen-deficient conditions inC. reinhardtii (Li et al., 2010b,c). Among all the macroele-ments tested, sulfur limitation promoted the highestaccumulation of starch and the longest maintenance timeof starch-enriched cells. Sulfur limitation was also success-fully tested under field conditions using the pilot-scaleoutdoor thin-layer solar photobioreactor (Branyikova et al.,2011).

However, in contrast to the strains above, a decrease inmacroelement concentration by depletion or omissionof nitrogen, phosphorus or sulfur from mineral medium(Figs. 2, 3, and 5) induced the hyper-accumulation (toabout 60% of DW) of reserve lipids instead of starch. Thisstress-induced high lipid production could, however, differ

Figure 3. Changes in biomass concentration (DW in g/L) (A–C) and relative starch and lipid content (% of DW) (D–F) in asynchronous cultures of P. kessleri aerated by air

(empty symbols) or by a mixture of air and CO2 (2%) (filled symbols). The cultures were grown in a laboratory photobioreactor under continuous illumination of an incident light

intensity of 780mmol/(m2 s) either in a complete mineral medium (1 medium) or in a medium of fivefold lower nitrogen content (0.2 N), or in 10-fold diluted medium (0.1 medium). The

initial biomass concentration (0.75 g/L) was identical for all variants.

102 Biotechnology and Bioengineering, Vol. 110, No. 1, January, 2013

substantially amongst various Chlorella and Parachlorellastrains (Pribyl et al., 2012). Nutrient limitation-inducedincreased lipid production has been described previouslymany times but hyper-accumulation to values of about 60%of DW has no often been achieved and in all cases, it wasattained at a much lower biomass density (Illman et al.,2000; Pribyl et al., 2012; Rodolfi et al., 2009).

Nutrient limitation with all tested macroelements had apositive impact on lipid hyper-accumulation. Previously,it was found that natural depletion of elements duringgrowth of C. vulgaris was the most effective way to increaselipid productivity (Stephenson et al., 2010). This approachalso effectively induced lipid accumulation, leading to veryhigh lipid productivity (0.325 g/L/day) (Pribyl et al., 2012).The problem is that exhaustion of elements to a levelenabling induction of lipid production is attained at so

high a biomass concentration that the mean light intensityper cell is too low to supply sufficient energy for lipidproduction. So, in order to use this very easy andeconomically advantageous procedure, it is necessary togrow cell cultures in an appropriately diluted medium.Depletion of nutrients in such a medium is attained at alower biomass concentration and thus at a higher mean lightintensity (see Fig. 6) which is needed for successful lipidproduction. Using this treatment, biomass accumulationceased (and oil production was triggered) at values between2 and 3.70 g/L of DW depending on the type of nutrientlimitation (Fig. 1B).

The commercial price of carbon dioxide, as a source ofcarbon for cellular metabolism in algal cultures, representsmore than 40% of the biomass price (Kadam, 1997).Utilization of waste CO2 from fuel gases of various sources

Figure 4. Time course of lipid accumulation during batch cultivation of P. kessleri in cultures aerated either by air or by a mixture of air and CO2 (2%). Lipid bodies were

stained using Nile Red (yellow); autofluorescence of chloroplasts is seen in red. Cultures grew either in a medium containing fivefold less nitrogen (0.2 N) or in fivefold diluted

medium (0.2 medium). The cultivation time for all variants is indicated in the left column. The same initial biomass concentration (0.3 g/L) was used for all cultures and no detectable

lipid bodies were seen in cells at the beginning (0 h; not shown in figure). Scale bar¼ 10mm.

Li et al.: Alga Parachlorella kessleri for Lipid Production 103

Biotechnology and Bioengineering

(incinerators, power stations, lime, biogas stations, etc.)proved to be feasible (Douskova et al., 2009).

In practice, however, it is sometimes difficult to locatebioreactors close to waste CO2 sources. However, a two-stepprocedure for lipid production can be applied because theexpensive components for algal growth (CO2 and fertilizers)are required only in the first step, for biomass accumulation.The second step, induction of lipid accumulation, can beperformed at a low CO2 concentration in air (Fig. 3E and F),due to slowed growth under nutrient limitation (Fig. 2); theinput of CO2 from air is sufficient.

For the large-scale production of lipid-enriched biomass,a relatively high input of light energy per algal cell must beassured. Under conditions of the present experiments,the minimummean light intensity for production of reservelipids was about 150mmol/(m2 s). This limit can differ invarious strains but to keep irradiation of cultures above thislimit is one of the most important requirements for lipidproduction.

In a thin layer algal suspension (7mm) grown in outdoorcultures, mean light intensity sufficient for lipid productioncan be attained at very high biomass concentrations (13 g/L)and relatively low incident light intensity (Fig. 7). Inaddition to the present findings, the convenience of a thinlayer scale-up (Doucha and Lıvansky, 2006, 2009) for biofuelproduction was verified, both for starch overproductionas a feedstock for bioethanol production (Branyikovaet al., 2011) and for overproduction of lipids for possibleproduction of biodiesel (Pribyl et al., 2012).

Starch is a major storage carbohydrate in many algae andhigher plants, and its biosynthesis shares commonprecursors with lipid biosynthesis. Previous studies haveindicated interactions between the two pathways, although

the regulation of carbon partitioning between starch andlipid biosynthetic pathways is still not well understood(Rawsthorne, 2002; Weselake et al., 2009).

Under nutrient limitation, some algal species or strainscan inhibit the starch biosynthetic pathway, redirecting the

Figure 5. Changes in total chlorophyll content in P. kessleri cultures aerated

either by air (empty symbols) or by a mixture of air and CO2 (2%) (filled symbols). The

cultures were grown in a laboratory photobioreactor under continuous illumination of

an incident light intensity of 780mmol/(m2 s) either in complete mineral medium

(1 medium) or in a medium containing fivefold less nitrogen (0.2 N), or in 10-fold

diluted medium (0.1 medium). The initial biomass concentration (0.75 g/L) was identical

for all.

Figure 6. Changes in biomass concentration (DW in g/L) (A) and in relative lipid

content (% of DW) (B) in cultures of P. kessleri grown at different mean light intensities

(C) as affected by various initial biomass concentrations (DW 4, 2, 1, 0.5, and 0.25 g/L,

curve 1, 2, 3, 4, 5 respectively) (A). The cultures were grown in a laboratory

photobioreactor at 308C at a constant incident light intensity of 780mmol/(m2 s) in

10-fold diluted complete mineral medium. Numerals on curves indicate the same

variants illustrated in panels (A–C).

104 Biotechnology and Bioengineering, Vol. 110, No. 1, January, 2013

photosynthetic carbon flux toward lipid biosynthesis,resulting in lipid hyper-accumulation as an alternate carbonand energy reserve for cells under stress (Li et al., 2011;Wang et al., 2009). After 2 days of nitrogen starvation, lipidbodies increased twofold in the C. reinhardtii starch-lessmutant compared to the wild type. The present findings inP. kessleri confirmed that this species, similarly to someother algae, accumulated lipids under conditions disruptingstarch accumulation. Our findings also suggest that thecessation of starch biosynthesis, as well as the slow decreasein starch content, was not related to lipid biosynthesis innutrient-limited cells, because a similar decrease in starchcontent also occurred in cultures grown in the completemedium with no or low accumulation of storage lipids.

In the green microalga Pseudochlorococcum sp., starchcontent decreased after nitrogen depletion, while neutrallipids rapidly increased to 52.1% of cell dry weight (Li et al.,2011). However, Li et al. (2011) found, in contrast to theresults of Wang et al. (2009) and the present findings, thatpartial inhibition of the enzymes of starch biosynthesis anddegradation resulted in a decrease in neutral lipid content,indicating that conversion of starch to neutral lipid mayhave contributed to the overall neutral lipid accumulation(Li et al., 2011). These contradictory results indicate thatdifferent species of algae may have evolved distinct strategiesto redirect carbon flux from carbohydrate pathways towardslipid biosynthesis. Some algae may accumulate similaramounts of starch and lipid under stress conditions suchas nitrogen limitation (Li et al., 2011; Ramazanov andRamazanov, 2006), whereas others only transiently accu-mulate starch (Collen et al., 2004).

To verify laboratory findings, experimentation usinglarge-scale production units is the only way to prove realoutcomes of any proposed approach. Unfortunately, to datealmost no large scale production studies giving relevant dataon the production of lipid-enriched algal biomass have beencarried out. There are only two exceptions; the studyon Nannochloropsis (Rodolfi et al., 2009) and the work onChlorella vulgaris (Pribyl et al., 2012). Though usingdifferent production systems, both studies demonstratedhigh maximal lipid productivity (over 0.250 g/L/day and upto 0.326 g/L/day, respectively), and total lipid content (60%and over 30% DW, respectively), albeit at relatively lowbiomass concentrations (up to 1.5 g DW/L and up to 6 gDW/L, respectively).

In the present experiments with P. kessleri grown in ascale-up thin layer outdoor PBR, if transferred into 10-fold

Table I. Biomass, lipid content, and productivity at different mean light intensities.

Biomass from

start to end (g/L)

Lipids ProductivityMean light intensity from

start to end mmol/(m2 s)Total (g/L) Abs. (g/L) Rel. % (DW) Biomass (g/L/d) Lipid (g/L/d)

4–7.8 3.8 1.2 15 0.80 0.34 125–69

2–4.75 2.75 1.1 25 0.42 0.30 160–104

1–3.4 2.4 1.6 44 0.40 0.60 183–125

0.5–1.75 1.25 1 45 0.30 0.50 216–148

0.25–1.5 1.25 1 51 0.35 0.58 292–160

Table II. Fatty acid composition of Parachlorella kessleri algal biomass in

cultures grown in the complete medium and nutrient-limited cultures.

Fatty acid Scheme

1 medium

(%)

0.2 N

(%)

0.2 medium

(%)

0.1 medium

(%)

Palmitic 16:0 32.4 24.9 28.8 25.7

Stearic 18:0 14.3 11.2 12.1 8.7

Oleic 18:1 8.4 18.5 16.8 10.1

Linoleic 18:2 42.5 42.0 40.0 52.4

Saturated/

unsaturated

0.917 0.597 0.874 0.550

Only FAs of an amount higher than 1% of the total FA content areshown.

Figure 7. Changes in biomass concentration in g/L (DW), cell volume in mm3 (V)

and cell number in 107/mL (N) (A), and relative content of both starch and lipid in %

of DW) (B) in cultures of P. kessleri grown in a scale-up thin-layer photobioreactor

in complete mineral medium for 14 days followed by addition of a 10-fold diluted

complete mineral medium. Variations in mean light intensity in mmol/(m2 s) are given in

the panel (B).

Li et al.: Alga Parachlorella kessleri for Lipid Production 105

Biotechnology and Bioengineering

diluted complete mineral medium, the lipid contentattained a maximum of 25% DW after 7 days, in a biomassof 14 g DW/L, corresponding to a lipid productivity of0.500 g/L/day. These achievements make this alga a viableoption for industrial lipid production. However, bottleneckswith low mean sunlight intensity must be prevented. It canbe attained by a combination of two conditions: Firstly anarea with sufficient sunlight intensity for the maximumnumber of sunny days per year and secondly a thin layerphotobioreactor, where turbulent flows of a thin layer ofalgae (<1 cm) increases the utilization of light energy. Thistype of PBR provides further benefits; a high algal densityreduces the risk of contamination by other species anddecreases the financial input for harvesting the biomass bycentrifugation.

References

Ballin G, Doucha J, Zachleder V, Setlık I. 1988. Macromolecular syntheses

and the course of cell cycle events in the chlorococcal alga Scenedesmus

quadricauda under nutrient starvation: Effect of nitrogen starvation.

Biol Plant 30:81–91.

Benemann JR. 1997. CO2 mitigation with microalgae systems. Energy

Convers Mgmt 38:S475–S479.

Biller P, Ross AB. 2011. Potential yields and properties of oil from the

hydrothermal liquefaction of microalgae with different biochemical

content. Bioresour Technol 102:215–225.

Branyikova I, Marsalkova B, Doucha J, Branyik T, Bisova K, Zachleder V,

Vıtova V. 2011. Microalgae––Novel highly-efficient starch producers.

Biotechnol Bioeng 108:766–776.

Brown LM. 1996. Uptake of carbon dioxide from flue gas by microalgae.

Energy Convers Mgmt 37(6–8):1363–1367.

Chen C-J, Yeh K-L, Aisyah R, Lee D-L, Chang J-C, 2011. Cultivation,

photobioreactor design and harvesting of microalgae for biodiesel

production: A critical review. Bioresour Technol 102:71–81.

Chisti Y. 2007. Biodiesel from microalgae. Biotechnol Adv 25:294–306.

Chisti Y, Yan J. 2011. Energy from algae: Current status and future trends.

Algal biofuels––A status report. Appl Energy 88:3277–3279.

Collen PN, Camitz A, Hancock RD, Viola R, Pedersen M. 2004. Effect of

nutrient deprivation and resupply on metabolites and enzymes related

to carbon allocation in Gracilaria tenuistipitata (Rhodophyta). J Phycol

40:305–314.

Deng X, Li Y, Fei X. 2009. Microalgae: A promising feedstock for biodiesel.

African J Microbiol Res 3:1008–1014.

Doucha J, Lıvansky K. 2006. Productivity, CO2/O2 exchange and hydraulics

in outdoor open high density microalgal (Chlorella sp.) photobior-

eactors operated in a Middle and Southern European climate. J Appl

Phycol 18(6):811–826.

Doucha J, Lıvansky K. 2009. Outdoor open thin-layer microalgal photo-

bioreactor: Potential productivity. J Appl Phycol 21:111–117.

Douskova I, Doucha J, Livansky K, Machat J, Novak P, Umysova D,

Zachleder V, Vitova M. 2009. Simultaneous flue gas bioremediation

and reduction of microalgal biomass production costs. Appl Microbiol

Biotechnol 82:179–185.

Eltgroth ML, Watwood RL, Wolfe GV. 2005. Production and cellular

localization of neutral long-chain lipids in the haprophyte algae Iso-

chorysis galbana and Emiliania huxleyi. J Phycol 41:1000–1009.

Francisco EC, Neves DB, Jacob-Lopes E, Franco TT. 2010. Microalgae as

feedstock for biodiesel production: Carbon dioxide sequestration, lipid

production, and biofuel quality. J Chem Technol Biotechnol 85(3):

395–403.

Goodson C, Roth R, Wang ZT, Goodenough U. 2011. Structural correlates

of cytoplasmic and chloroplast lipid body synthesis in Chlamydomonas

reinhardtii and stimulation of lipid body production with acetate boost.

Eukaryot Cell 10:1592–1606.

Halim R, Gladman B, Danquah MK, Webley PA. 2011. Oil extraction from

microalgae for biodiesel production. Bioresour Technol 102:130–134.

Hu Q, SommerfeldM, Jarvis E, Ghirardi M, Posewitz M, Seibert M, Darzins

A. 2008. Microalgal triacylglycerols as feedstocks for biofuel produc-

tion: Perspectives and advances. Plant J 54:621–639.

Illman AM, Scragg AH, Shales SW. 2000. Increase in Chlorella strains

calorific values when grown in low nitrogen medium. Enzym Microb

Technol 27:631–635.

Kadam KL. 1997. Plant flue gas as a source of CO2 for microalgae

cultivation. Economic impact of different process options. Energy

Convers Mgmt 38:S505–S510.

Krienitz L, Hegewald EH, Hepperle D, Huss VAR, Rohr T, Wolf M. 2004.

Phylogenetic relationship of Chlorella and Parachlorella gen nov.

(Chlorophyta, Trebouxiophyceae). Phycologia 43:529–542.

Lee DH. 2011. Algal biodiesel economy and competition among bio-fuels.

Bioresour Technol 102:43–49.

Li Y, Han D, Hu G, Dauvillee D, Sommerfeld M, Ball S, Hu Q. 2010a.

Chlamydomonas starchless mutant defective in ADP-glucose pyropho-

sphorylase hyper-accumulates triacylglycerol. Metab Eng 12:387–391.

Li Y, Han D, Hu G, Dauvillee D, Sommerfeld M, Ball S, Hu Q. 2010b.

Chlamydomonas starchless mutant defective in ADP-glucose pyropho-

sphorylase hyper-accumulates triacylglycerol. Metab Eng 12:387–391.

Li Y, Han D, Hu G, Sommerfeld M, Hu Q. 2010c. Inhibition of starch

synthesis results in overproduction of lipids in Chlamydomonas rein-

hardtii. Biotechnol Bioeng 107:258–268.

Li Y, Han D, Sommerfeld M, Hu Q. 2011. Photosynthetic carbon partition-

ing and lipid production in the oleaginous microalga Pseudochloro-

coccum sp. (Chlorophyceae) under nitrogen-limited conditions.

Bioresour Technol 102.

MacKinney G. 1941. Absorption of light by chlorophyll solutions. J Biol

Chem 140:315–322.

McCready RM, Guggolz J, Silviera V, Owens HS. 1950. Determination of

starch and amylose in vegetables. Analyt Chem 22:1156–1158.

Mutlu YB, Isık O, Uslu L, Koc K, Durmaz Y. 2011. The effects of nitrogen

and phosphorus deficiencies and nitrite addition on the lipid content

of Chlorella vulgaris (Chlorophyceae). African J Biotechnol 10:453–

456.

Patil PD, Gude VG,Mannarswamy A, Deng S, Cooke P, McGee SM, Rhodes

I, Lammers P, Nirmalakhandan N. 2011. Optimization of direct

conversion of wet algae to biodiesel under supercritical methanol

conditions. Bioresour Technol 102:118–122.

Pribyl P, Cepak V, Zachleder V. 2012. Production of lipids in 10 strains of

Chlorella and Parachlorella, and enhanced lipid productivity in Chlo-

rella vulgaris. Appl Microbiol Biotechnol 94:549–561.

Ramazanov A, Ramazanov Z. 2006. Isolation and characterization of a

starchless mutant of Chlorella pyrenoidosa STL-PI with a high growth

rate, and high protein and polyunsaturated fatty acid content. Phycol

Res 54:255–259.

Rawsthorne S. 2002. Carbon flux and fatty acid synthesis in plants. Prog

Lipid Res 41:182–196.

Reitan KI, Rainuzzo JR, Olsen Y. 1994. Effect of nutrient limitation on fatty

acid and lipid content of marine microalgae. J Phycol 30:972–979.

Rezanka T, Petrankova M, Cepak V, Sigler K, Cajthaml T. 2010. Trachy-

discus minutus, a new biotechnological source of eicosapentaenoic acid.

Folia Microbiol 55:265–269.

Rodolfi L, Zittelli GC, Bassi N, Padovani G, Biondi N, Bonini G, Tredici

MR. 2009. Microalgae for oil: Strain selection, induction of lipid

synthesis and outdoor mass cultivation in a low-cost photobioreactor.

Biotechnol Bioeng 102(1):100–112.

Setlık I, Ballin G, Doucha J, Zachleder V. 1988. Macromolecular syntheses

and the course of cell cycle events in the chlorococcal alga Scenedesmus

quadricauda under nutrient starvation: Effect of sulphur starvation.

Biol Plant 30:161–169.

Sheehan J, Dunahay T, Benemann J, Roessler PG. 1998. US Department of

Energy’s Office of Fuels Development, July 1998. A Look Back at the

US, Department of Energy’s Aquatic Species Program—Biodiesel from

106 Biotechnology and Bioengineering, Vol. 110, No. 1, January, 2013

Algae, Close Out Report TP-580-24190. Golden, CO: National Renew-

able Energy Laboratory.

Singh A, Nigam PS, Murphy JD. 2011a. Mechanism and challenges in

commercialisation of algal biofuels. Bioresour Technol 102:26–34.

Singh A, Nigam PS, Murphy JD. 2011b. Renewable fuels from algae:

An answer to debatable land based fuels. Bioresource Technol

102:10–16.

Stephenson AI, Dennis JS, Howe CJ, Scott SA, Smith AG. 2010. Influence of

nitrogen-limitation regime on the production by Chlorella vulgaris of

lipids for biodiesel feedstocks. Biofuels 1:47–58.

Tornabene TG, Holzer G, Lien S, Burris N. 1983. Lipid composition of the

nitrogen starved green alga Neochloris oleoabundans. Enzyme Microb

Technol 5:435–440.

Wang ZT, Ullrich N, Joo S, Waffenschmidt S, Goodenough U. 2009. Algal

lipid bodies: Stress induction, purification, and biochemical character-

ization in wild-type and starchless Chlamydomonas reinhardtii. Eukar-

yot Cell 8(12):1856–1868.

Weselake RJ, Taylor DC, Rahman MH, Shah S, Laroche A, McVetty PBE,

Harwood JL. 2009. Increasing the flow of carbon into seed oil.

Biotechnol Adv 27:866–878.

Yao C, Ai J, Cao X, Xue S, ZhangW. 2012. Enhancing starch production of a

marine green microalga Tetraselmis subcordiformis through nutrient

limitation. Bioresour Technol 118:438–444.

Yu Y, Chen BL, You WL. 2007. Identification of the alga known as

Nannochloropsis Z-1 isolated from a prawn farm in Hainan, China

as Chlorella. World J Microbiol Biotechnol 23:207–210.

Zachleder V, Ballin G, Doucha J, Setlık I. 1988. Macromolecular syntheses

and the course of cell cycle events in the chlorococcal alga Scenedesmus

quadricauda under nutrient starvation: Effect of phosphorus starvation.

Biol Plant 30:92–99.

Li et al.: Alga Parachlorella kessleri for Lipid Production 107

Biotechnology and Bioengineering