Open Access

Growth of Chlamydomonas reinhardtii in acetate-free medium when co-cultured with alginate-encapsulated, acetate-producing strains of Synechococcus sp. PCC 7002

  • Jesse B Therien1,
  • Oleg A Zadvornyy1,
  • Matthew C Posewitz3,
  • Donald A Bryant1, 2 and
  • John W Peters1Email author
Biotechnology for Biofuels20147:154

DOI: 10.1186/s13068-014-0154-2

Received: 5 June 2014

Accepted: 2 October 2014

Published: 18 October 2014

Abstract

Background

The model alga Chlamydomonas reinhardtii requires acetate as a co-substrate for optimal production of lipids, and the addition of acetate to culture media has practical and economic implications for algal biofuel production. Here we demonstrate the growth of C. reinhardtii on acetate provided by mutant strains of the cyanobacterium Synechococcus sp. PCC 7002.

Results

Optimal growth conditions for co-cultivation of C. reinhardtii with wild-type and mutant strains of Synechococcus sp. 7002 were established. In co-culture, acetate produced by a glycogen synthase knockout mutant of Synechococcus sp. PCC 7002 was able to support the growth of a lipid-accumulating mutant strain of C. reinhardtii defective in starch production. Encapsulation of Synechococcus sp. PCC 7002 using an alginate matrix was successfully employed in co-cultures to limit growth and maintain the stability.

Conclusions

The ability of immobilized strains of the cyanobacterium Synechococcus sp. PCC 7002 to produce acetate at a level adequate to support the growth of lipid-accumulating strains of C. reinhartdii offers a potentially practical, photosynthetic alternative to providing exogenous acetate into growth media.

Keywords

Biofuels Algae Cyanobacteria Lipid production Acetate production Co-culture Alginate immobilization

Background

With the ongoing increase in global energy demand, the development of alternative energy sources has been at the forefront of recent modern research. Of the various alternative energies, bioenergy has been of interest because of its many potential benefits over the currently used petroleum, natural gas, and coal. These benefits include carbon neutrality, renewability, low environmental toxicity, and reduction of dependence on foreign energy sources. Biodiesel and bioethanol, which together currently account for the major proportion of biofuels, are mainly produced from higher plants [1]. These fuels can replace diesel and gasoline, respectively, in conventional engines without modification. Of these two biofuels, biodiesel has been considered the most viable option in the United States [2]. However, the production of biodiesel from crops grown on arable land is controversial due to the potential implications of the competition with agricultural food production [3],[4]. One alternative to using food crops as oil sources is to produce biofuels using microalgae and/or cyanobacteria, which can be grown in areas not suitable for crop growth, such as deserts [5],[6]. Microalgae can be used to assimilate CO2 and produce multiple fuel molecules, including hydrogen, starch, and lipids [7]-[9]. Starch can be fermented to hydrogen or ethanol, while lipids can be converted to biodiesel [10]-[12].

Some lipid-accumulating microalgae, such as Chlamydomonas reinhardtii, are able to grow photoautotrophically on sunlight and CO2, chemotrophically on acetate, or photomixotrophically in a combination of these two growth modes [13],[14]. Optimal lipid production by C. reinhardtii is observed under photomixotrophic conditions in the presence of acetate, which could present economic and practical challenges for large-scale production of algal-based biofuels [15],[16]. An alternative is to co-culture lipid-producing C. reinhardtii with an acetate-producing cyanobacterium, such as certain Synechococcus sp., that can naturally produce acetate during photosynthesis and/or fermentation [17]-[19].

In this study, we present results of co-culturing wild-type and a lipid-accumulating (sta6) strain of C. reinhardtii, and an acetate-accumulating strain (glgA1) of Synechococcus sp. PCC 7002. We demonstrate sustained photomixotrophic growth of C. reinhardtii with acetate produced by Synechococcus 7002.

Results and discussion

Effect of temperature on growth of Synechococcus sp. PCC 7002 and C. reinhardtii

To determine conditions under which both organisms could grow well together, the wild-type and mutant strains of C. reinhardtii and Synechococcus sp. PCC 7002 were grown individually on Tris-Acetate-Phosphate (TAP) medium, a standard medium for C. reinhardtii, and A+ medium, a standard medium for Synechococcus sp. PCC 7002. Consistent with previous reports, the results of our preliminary experiments showed that the optimum temperature for the growth of C. reinhardtii is ~30°C [20]-[22] and that for Synechococcus 7002 ~ 38°C [23]. An increase in temperature from 30°C to 34°C and then to 38°C increased the lag phase in the C. reinhardtii cultures (Figure 1A). In contrast, decreasing the temperature from 38°C to 34°C and finally to 30°C did not significantly affect the growth of either wild-type (Figure 1B) or mutant strains of Synechococcus sp. PCC 7002 under the light intensities and CO2 concentrations used in this study (data not shown). Therefore subsequent co-culture optimizations were conducted at 30°C.
Figure 1

Effect of temperature on growth of C. reinhartdii and Synechococcus sp . PCC 7002. Cultures of C. reinhartdii (A) and Synechococcus sp. PCC 7002 (B) were grown at 30°C (circles), 34°C (squares), and 38°C (triangles).

Effect of media composition on growth of C. reinhardtii and Synechococcus sp. PCC 7002

The main difference between TAP medium (C. reinhardtii) and A+ medium (Synechococcus sp. PCC 7002) is that medium A+ is a marine medium that contains 300 mM sodium chloride. Other differences include the use of ammonium chloride as the nitrogen source in TAP medium versus sodium nitrate in A+ medium, the presence of acetate in TAP, and the presence of vitamin B12 in A+ medium. To accommodate the growth of both C. reinhardtii and Synechococcus sp. PCC 7002 in co-culture, both the TAP and A+ media were modified (henceforth referred to as modified TAP and modified A+, respectively). Modified TAP medium consists of TAP medium supplemented with sodium nitrate and vitamin B12; modified A+ medium consists of standard A+ medium supplemented with ammonium chloride and acetic acid, with the sodium chloride concentration reduced to 150 mM. (Additional file 1: Table S1). The results of our experiments showed that C. reinhardtii cultures are unable to grow on A+ medium and/or modified A+ medium at 30°C (Figure 2, and insert in Figure 2) and 38°C (data not shown). However, Synechococcus sp. PCC 7002 was able to grow on both TAP and modified TAP media at 30°C (Figure 2).
Figure 2

Growth of C. reinhartdii and Synechococcus sp. PCC 7002 on different types of media. Cultures of C. reinhartdii (black bars) and Synechococcus sp. PCC 7002 (gray bars) were grown on TAP, co-culture media (TAP mod), A+, modified A+ media at 30°C after 124 hours. The difference in the media composition is described in the text and in the Methods section.

Effect of acetate on growth of C. reinhardtii and Synechococcus sp. PCC 7002

To investigate the effects of acetate on the growth of both wild-type and mutant cultures of C. reinhardtii and Synechococcus sp. PCC 7002, cultures were grown on TAP medium containing acetate and on a modified form of TAP medium that lacked acetate (termed co-culture medium). The results of these experiments showed that the growth rates of wild-type and mutant cultures of C. reinhartdii were higher in the presence of acetate when compared to cultures grown in the absence of acetate. Cultures of C. reinhardtii grown in acetate-free medium showed little growth under these conditions (Figure 3A). Although Synechococcus sp. PCC 7002 cultures grown without acetate had slightly lower final cell densities than cultures grown in the presence of acetate, the growth rates with and without acetate were similar for both wild-type and mutant strains (Figure 3B). During the growth of wild-type and sta6 mutant strains of C. reinhardtii, acetate consumption correlated with an increase in cell number (Figure 4A). Acetate production by the glgA1 mutant of Synechococcus 7002 was higher than that of the wild type (Figure 4B).
Figure 3

Effect of acetate on growth of C. reinhartdii and Synechococcus sp. PCC 7002. The cultures of C. reinhartdii (A) and Synechococcus sp. PCC 7002 (B) were grown in the presence (closed symbols) and absence (open symbols) of acetate. Both C. reinhartdii and Synechococcus sp. PCC 7002 wild-type cultures are shown in circles. The sta6 mutant of C. reinhartdii and the glgA mutant of Synechococcus sp. PCC 7002 are shown in squares.

Figure 4

Growth and consumption of acetate by C. reinhartdii cultures, and growth and production of acetate by Synechococcus sp. PCC 7002 cultures. The cultures of wild-type (closed circles) and sta6 mutant (closed squares) of C. reinhartdii (A) and wild-type (closed circles) and glgA (closed squares) of Synechococcus sp. PCC 7002 (B) grown on co-culture media at pH 7.0 and 30°C. Acetate consumption (A) by wild-type (open circles) and sta6 mutant (open squares) C. reinhartdii and production (B) by wild-type (open circles) and glgA mutant (open squares) Synechococcus sp. PCC 7002 were measured during the cell growth.

Optimizing co-cultures

Based on the initial experiments, the sta6 mutant of C. reinhardtii was grown on co-culture medium at pH 7.0 and 30°C with the glgA1 mutant of Synechococcus sp. PCC 7002. To investigate the effect of initial cell number in the inoculum during co-culturing of free cells, different initial ratios, 1:1 and 1:10, of C. reinhardtii to Synechococcus sp. PCC 7002 cells were tested. The ratio 1:10 was chosen because Synechococcus sp. PCC 7002 appeared to produce less acetate than required to produce optimal growth of C. reinhardtii, and it was rationalized that a larger number of Synechococcus sp. PCC 7002 cells might be advantageous in the co-cultures. However, the results of the co-culturing experiments with free cells revealed that Synechococcus sp. PCC 7002 cells grew faster than C. reinhardtii at both initial cell ratios and eventually took over the co-culture (data not shown). To avoid the problem of overgrowth by Synechococcus sp. PCC 7002, the cells were encapsulated in alginate beads to slow their growth (see Methods section for details). The growth of the C. reinhardtii sta6 mutant in the presence of control (empty) beads in modified TAP media showed that the alginate encapsulating matrix had no effect on the growth of C. reinhardtii cells (data not shown). The results of the experiments using alginate-encapsulated Synechococcus sp. PCC 7002 cells showed that the growth of the lipid-accumulating C. reinhardtii sta6 strain could be supported by the presence of the acetate-producing Synechococcus sp. PCC 7002 glgA1 strain (Figure 5) and that immobilization controlled the growth of Synechococcus sp. PCC 7002 and kept the co-cultures from being overgrown.
Figure 5

Co-culturing C. reinhartdii and Synechococcus sp. PCC 7002. The C. reinhartdii sta6 mutant was grown with alginate bead immobilized Synechococcus sp. PCC 7002 wild-type cells (blue squares), glgA1 cells (green diamonds), and with empty beads (red triangles) in co-culture on modified TAP media without acetate at pH 7.0. Growth of Synechococcus sp. PCC 7002 cells leaking out of the beads is shown in gray squares. Acetate production by alginate bead immobilized Synechococcus sp. PCC 7002 glgA1 mono-culture is shown in stars. The inset shows alginate beads of Synechococcus sp. (left) and an empty bead (right).

Conclusions

We have developed media formulations and growth conditions that support co-cultures of wild-type and the acetate-producing glgA1 strain of the Synechococcus sp. PCC 7002 and wild-type and lipid-accumulating sta6 mutant of C. reinhardtii (Additional file 2: Figure S1). A temperature of 30°C supported the growth of both organisms; however, Synechococcus sp. PCC 7002 overtook the culture when both organisms were present as free cells. Alginate encapsulation was found to be an effective way to slow the growth of Synechococcus sp. PCC 7002 while at the same time allowing the production of sufficient acetate to stimulate growth and lipid storage of C. reinhardtii. The encapsulation of Synechococcus sp. PCC 7002 also provides a potential mechanism for separating and harvesting C. reinhardtii for lipids and recycling of the Synechococcus sp. PCC 7002 cells. The results reported here provide a strong proof of concept for supplanting the provision of costly medium components through co-culturing that can be optimized through metabolic engineering and implementation of enhanced acetate-producing cyanobacteria.

Methods

Growth of microorganisms

Inoculum cultures of both Synechococcus sp. PCC 7002 and C. reinhardtii (25 ml) were prepared in standard growth media for each organism in 50-ml Erlenmeyer flasks oscillating at 100 rpm at a constant illumination of 100 μmol photons m-2-s-1 with warm white fluorescent light. Wild-type and glgA1 knockout mutant strains of Synechococcus sp. PCC 7002 were grown in medium A+ (pH = 8.0) [24] (Additional file 1: Table S1) and were incubated at 38°C while sparging with 1% v/v CO2 in air. Wild-type C. reinhardtii and sta6 mutant strains were grown in TAP media (pH = 7.0) incubated at 30°C. The optical density was monitored at 750 nm (ThermoSpectronic Bio Mate 3 spectrophotometer) for Synechococcus sp. PCC 7002 and C. reinhardtii strains. Cultures were harvested by centrifugation for 10 min at 8,000 × g, washed in the medium to be used, centrifuged again, and finally resuspended in the appropriate medium and used to inoculate different culture conditions for co-culturing. Optimal co-culturing conditions were determined by first growing wild-type Synechococcus sp. PCC 7002 and C. reinhardtii strains individually at 30°C, 34°C, and 38°C on TAP medium (standard C. reinhardtii growth medium) or A+ medium (standard Synechococcus sp. PCC 7002 growth medium). The formulation of the TAP media, A+ medium, and modified derivatives of each can be found in the supplemental data (Additional file 1: Table S1). For the study, standard TAP medium was modified by eliminating acetic acid and supplementing with 1 g NaNO3 L-1 and 4 μg vitamin B12 L-1, and this formulation was termed co-culture medium. A+ medium was supplemented with 7.5 g NH4Cl L-1 and 1 ml glacial acetic acid L-1, and the concentration of NaNO3 and NaCl was reduced by 50%. Co-cultures were inoculated with a mixture of seed cultures grown in the appropriate culture media, and the optical densities were monitored at both 600 nm and 750 nm. The cultures were monitored for bacterial contamination by microscopic examination, and cell numbers were determined by direct cell counting with a hemocytometer.

Cell immobilization

Synechococcus sp. PCC 7002 cultures were harvested at an OD650 nm ranging from 0.8-1.0 (ThermoSpectronic Bio Mate 3 spectrophotometer) via centrifugation and resuspended in 1/5 the original media volume, yielding a concentrated cell suspension. Alginate beads were made by dissolving 3 g of sodium alginate directly in 100 ml of concentrated culture media and adding it dropwise with a syringe and needle into a 1% (w/v) solution of CaCl2 in growth media from a height of at least 30 cm [25]. The bead size was controlled by needle gauge: 22 G yields beads approximately 2 mM in diameter and 18 G yields beads of 3 mM, while 4-mm beads are made directly from a 10-ml syringe without a needle. Beads were formed immediately and were allowed to harden further in the CaCl2 solution for 15 min. After hardening, the beads were removed from the CaCl2 solution and added to fresh growth media. To prevent leakage of cells, a cell-free layer of alginate was used to coat the beads. To produce the coating, the beads were submerged in a 3% (w/v) alginate solution in growth medium followed by transfer to a 1% (w/v) solution of CaCl2 in growth medium to allow hardening of the cell-free alginate layer.

Acetate determination

Acetate concentrations were determined by 1H-NMR [26]. Samples were centrifuged for 10 min at 14,000× g and 500 μl of the supernatant was transferred to an NMR tube to which 50 μl of D2O was added. Spectra were collected on a Bruker DRX500 NMR spectrometer operating at a frequency of 500.13 MHz with a 1D NOESY pulse sequence and a 100-ms mixing time at 32 K. Typically, 64 4.28-s scans were collected for each sample with a pre-saturation pulse to suppress the water signal.

Authors’ contributions

JBT and OAZ contributed equally in conducting the experiments. JWP, DAB, MCP, OAZ, and JBT designed the experiments, analyzed the data, and wrote the article. All authors read and approved the final manuscript.

Additional files

Declarations

Acknowledgements

The authors would like to thank Dr. Scott Busse, Interim Director of Montana State University NMR facility, for assistance in measuring the acetate concentrations. The project was supported by a grant from DOE Energy Efficiency and Renewable Energy (DE-EE0003136) to JWP and DAB, and AFOSR (FA9550-14-1-0147) to MCP and JWP.

Authors’ Affiliations

(1)
Department of Chemistry and Biochemistry, Montana State University
(2)
Department of Biochemistry and Molecular Biology, The Pennsylvania State University
(3)
Department of Chemistry and Geochemistry, Colorado School of Mines

References

  1. Ren JZ, Manzardo A, Mazzi A, Fedele A, Scipioni A: Emergy analysis and sustainability efficiency analysis of different crop-based biodiesel in life cycle perspective. Scientific World J 2013, 2013: 1-12.Google Scholar
  2. Posten C, Schaub G: Microalgae and terrestrial biomass as source for fuels - a process view. J Biotechnol 2009, 142: 64-69. 10.1016/j.jbiotec.2009.03.015View ArticleGoogle Scholar
  3. Schenk PM, Thomas-Hall SR, Stephens E, Marx UC, Mussgnug JH, Posten C, Kruse O, Hankamer B: Second generation biofuels: high-efficiency microalgae for biodiesel production. Bioenergy Res 2008, 1: 20-43. 10.1007/s12155-008-9008-8View ArticleGoogle Scholar
  4. Dwidar M, Kim S, Jeon BS, Um Y, Mitchell RJ, Sang BI: Co-culturing a novel Bacillus strain with Clostridium tyrobutyricum ATCC 25755 to produce butyric acid from sucrose. Biotechnol Biofuels 2013, 6: 35. 10.1186/1754-6834-6-35View ArticleGoogle Scholar
  5. Hannon M, Gimpel J, Tran M, Rasala B, Mayfield S: Biofuels from algae: challenges and potential. Biofuels 2010, 1(5):763-784. 10.4155/bfs.10.44View ArticleGoogle Scholar
  6. Campbell PK, Beer T, Batten D: Life cycle assessment of biodiesel production from microalgae in ponds. Bioresour Technol 2011, 102: 50-56. 10.1016/j.biortech.2010.06.048View ArticleGoogle Scholar
  7. Beer LL, Boyd ES, Peters JW, Posewitz MC: Engineering algae for biohydrogen and biofuel production. Curr Opin Biotechnol 2009, 20: 264-271. 10.1016/j.copbio.2009.06.002View ArticleGoogle Scholar
  8. Radakovits R, Eduafo PM, Posewitz MC: Genetic engineering of fatty acid chain length in Phaeodactylum tricornutum . Metab Eng 2013, 13: 89-95. 10.1016/j.ymben.2010.10.003View ArticleGoogle Scholar
  9. Lakaniemi AM, Hulatt CJ, Thomas DN, Tuovinen OH, Puhakka JA: Biogenic hydrogen and methane production from Chlorella vulgaris and Dunaliella tertiolecta biomass. Biotechnol Biofuels 2011, 4: 34. 10.1186/1754-6834-4-34View ArticleGoogle Scholar
  10. Wahlen BD, Willis RM, Seefeldt LC: Biodiesel production by simultaneous extraction and conversion of total lipids from microalgae, cyanobacteria, and wild mixed-cultures. Bioresour Technol 2011, 102: 2724-2730. 10.1016/j.biortech.2010.11.026View ArticleGoogle Scholar
  11. Wang JF, Li RM, Lu D, Ma S, Yan YP, Li WJ: A quick isolation method for mutants with high lipid yield in oleaginous yeast. World J Microbiol Biotechnol 2009, 25: 921-925. 10.1007/s11274-009-9960-2View ArticleGoogle Scholar
  12. Meuser JE, Ananyev G, Wittig LE, Kosourov S, Ghirardi ML, Seibert M, Dismukes GC, Posewitz MC: Phenotypic diversity of hydrogen production in chlorophycean algae reflects distinct anaerobic metabolisms. J Biotechnol 2009, 142: 21-30. 10.1016/j.jbiotec.2009.01.015View ArticleGoogle Scholar
  13. Stern DB, Witman GB: The Chlamydomonas Sourcebook: Organellar and Metabolic Processes. Academic Press, Oxford UK; 1989.Google Scholar
  14. Zabawinski C, Van den Koornhuyse N, D’Hulst C, Schlichting R, Giersch C, Delrue B, Lacroix JM, Preiss J, Ball S: Starchless mutants of Chlamydomonas reinhardtii lack the small subunit of a heterotetrameric ADP-glucose pyrophosphorylase. J Bacteriol 2001, 183: 1069-1077. 10.1128/JB.183.3.1069-1077.2001View ArticleGoogle Scholar
  15. Work VH, Radakovits R, Jinkerson RE, Meuser JE, Elliott LG, Vinyard DJ, Laurens LM, Dismukes GC, Posewitz MC: Increased lipid accumulation in the Chlamydomonas reinhardtii sta7-10 starchless isoamylase mutant and increased carbohydrate synthesis in complemented strains. Eukaryot Cell 2010, 9: 1251-1261. 10.1128/EC.00075-10View ArticleGoogle Scholar
  16. Sager R, Granick S: Nutritional studies with Chlamydomanas reinhardtii . Ann N Y Acad Sci 1953, 56: 831-838. 10.1111/j.1749-6632.1953.tb30261.xView ArticleGoogle Scholar
  17. McNeely K, Xu Y, Bennette N, Bryant DA, Dismukes GC: Redirecting reductant flux into hydrogen production via metabolic engineering of fermentative carbon metabolism in a cyanobacterium. Appl Environ Microbiol 2010, 76: 5032-5038. 10.1128/AEM.00862-10View ArticleGoogle Scholar
  18. Xu Y, Guerra LT, Li ZK, Ludwig M, Dismukes GC, Bryant DA: Altered carbohydrate metabolism in glycogen synthase mutants of Synechococcus sp strain PCC 7002: Cell factories for soluble sugars. Metab Eng 2013, 16: 56-67. 10.1016/j.ymben.2012.12.002View ArticleGoogle Scholar
  19. Guerra LT, Levitan O, Frada MJ, Sun JS, Falkowski PG, Dismukes GC: Regulatory branch points affecting protein and lipid biosynthesis in the diatom Phaeodactylum tricornutum . Biomass Bioenergy 2013, 59: 306-315. 10.1016/j.biombioe.2013.10.007View ArticleGoogle Scholar
  20. Harris EH: The Chlamydomonas Sourcebook: Introduction to Chlamydomonas and Its Laboratory Use. Academic Press, Oxford UK; 1989.Google Scholar
  21. Tsygankov AA, Kosourov SN, Tolstygina IV, Ghirardi ML, Seibert M: Hydrogen production by sulfur-deprived Chlamydomonas reinhardtii under photoautotrophic conditions. Int J Hydrogen Energy 2006, 31: 1574-1584. 10.1016/j.ijhydene.2006.06.024View ArticleGoogle Scholar
  22. Laurinavichene TV, Fedorov AS, Ghirardi ML, Seibert M, Tsygankov AA: Demonstration of sustained hydrogen photoproduction by immobilized, sulfur-deprived Chlamydomonas reinhardtii cells. Int J Hydrogen Energy 2006, 31: 659-667. 10.1016/j.ijhydene.2005.05.002View ArticleGoogle Scholar
  23. Ludwig M, Bryant DA: Synechococcus sp. strain PCC 7002 transcriptome: acclimation to temperature, salinity, oxidative stress, and mixotrophic growth conditions. Front Microbiol 2012, 3: 354.Google Scholar
  24. Stevens SE, Myers J: The production of hydrogen peroxide by blue-green algae - a survey. J Phycol 1973, 9: 427-430.Google Scholar
  25. Eldridge D: A novel approach to photosynthesis practicals. School Sci Rev 2004, 85: 37-45.Google Scholar
  26. Rajabzadeh M: Determination of unknown concentrations of sodium acetate using the method of standard addition and proton NMR: an experiment for the undergraduate analytical chemistry laboratory. J Chem Educ 2012, 89: 1454-1457. 10.1021/ed200577vView ArticleGoogle Scholar

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© Therien et al.; licensee BioMed Central Ltd. 2014

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