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Proposal full title: Algae and aquatic biomass for a sustainable production of 2nd generation biofuels Proposal acronym: AquaFUELs Type of funding scheme: Cooperation Theme 5 Energy Taxonomy, Biology and Biotechnology Name of the coordinating person: Dr. Raffaello Garofalo Coordinator email: ebb@ebb‐eu.org Coordinator phone: +32 2 7632477 Coordinator fax: +32 2 7630457 REV
Date
Organisation
Beneficiaries involved
FINAL
20/05/2011
Natascia Biondi, Mario Tredici
UNIFI
UNIFI
Dissemination level PU
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Table of contents
INTRODUCTION ................................................................................................................................................. 5 1 1.1 IMPORTANCE OF ALGAE AND AQUATIC BIOMASS FOR BIOFUELS ......................................................................... 6 1.1.1 Suitability of algae as biomass producers ..................................................................................................... 6 1.1.2 Sustainability, the strategic advantage of algal biofuels ............................................................................... 6 1.2 RATIONALE OF THE DOCUMENT ......................................................................................................................... 8 1.3 TARGET GROUPS ................................................................................................................................................ 8 1.4 PROBLEMS INCURRED ........................................................................................................................................ 9 1.5 COMMON ERRONEOUS "MYTH" .......................................................................................................................... 9 2 CRITERIA FOR STRAIN SELECTION ............................................................................................................ 10 2.1 PRODUCTIVITY ................................................................................................................................................. 10 2.2 ROBUSTNESS .................................................................................................................................................... 10 2.3 HARVESTABILITY ............................................................................................................................................. 10 2.4 BIOMASS COMPOSITION ................................................................................................................................... 10 2.5 PROCESSABILITY / EXTRACTABILITY ................................................................................................................ 11 2.6 ADDED VALUE OF CO-PRODUCTS ..................................................................................................................... 11 2.7 LOCAL ORIGIN OF STRAINS ............................................................................................................................... 11 3 BIOLOGY OF ALGAE....................................................................................................................................... 12 3.1 CYANOBACTERIA ............................................................................................................................................. 12 3.2 CHLOROPHYTA (GREEN ALGAE)...................................................................................................................... 15 3.3 RHODOPHYTA (RED ALGAE)............................................................................................................................ 17 3.4 HETEROKONTOPHYTA ...................................................................................................................................... 18 3.4.1 Phaeophyceae (Brown algae)...................................................................................................................... 19 3.4.2 Eustigmatophyceae ..................................................................................................................................... 21 3.4.3 Other classes ............................................................................................................................................... 21 3.5 LABYRINTHULEA (PHYLUM HETEROKONTA) ................................................................................................... 21 3.6 BACILLARIOPHYTA (DIATOMS)........................................................................................................................ 22 3.7 HAPTOPHYTA ................................................................................................................................................... 24 3.8 DINOPHYTA (DINOFLAGELLATES) ................................................................................................................... 24 3.9 OTHER ALGAL GROUPS..................................................................................................................................... 26 4 BIOTECHNOLOGY OF ALGAE ...................................................................................................................... 28 4.1 INTRODUCTION ................................................................................................................................................ 28 4.2 CULTIVATION SYSTEMS ................................................................................................................................... 29 4.2.1 Open ponds ................................................................................................................................................. 30 4.2.2 Photobioreactors ......................................................................................................................................... 31 Main photobioreactors designs ...............................................................................................................................................32 Polyethylene bags and vertical columns.............................................................................................................................32 Tubular PBR ......................................................................................................................................................................33 Flat photobioreactors (panels) ............................................................................................................................................35
4.2.3 Sustainability of different cultivation systems............................................................................................ 35 4.3 HARVESTING METHODS.................................................................................................................................... 36 4.4 BIOTECHNOLOGY OF THE MAJOR MICROALGAL GROUPS .................................................................................. 37 4.4.1 Cyanobacteria ............................................................................................................................................. 37 4.4.2 Chlorophyta (Green Algae)......................................................................................................................... 38 4.4.3 Rhodophyta (Red Algae) ............................................................................................................................ 39 4.4.4 Heterokontophyta........................................................................................................................................ 39 4.4.5 Labyrinthulea (phylum Heterokonta).......................................................................................................... 40 4.4.6 Bacillariophyta (Diatoms)........................................................................................................................... 40 4.4.7 Haptophyta.................................................................................................................................................. 41 4.4.8 Dinophyta (Dinoflagellates)........................................................................................................................ 41 4.5 BIOTECHNOLOGY AND USES FOR MACROALGAE .............................................................................................. 41 4.6 BIOTECHNOLOGY OF OTHER AQUATIC BIOMASS ............................................................................................. 43 5 SYMBOLOGY.................................................................................................................................................... 44 6 REFERENCES .................................................................................................................................................... 45 7 PROKARYOTIC MICROALGAE ..................................................................................................................... 51 7.1 CYANOBACTERIA ............................................................................................................................................. 51 7.1.1 Arthrospira sp. (common name spirulina) .................................................................................................. 51 7.1.2 Phormidium sp............................................................................................................................................ 54 7.1.3 Anabaena sp................................................................................................................................................ 57 7.1.4 Synechococcus sp........................................................................................................................................ 60 7.1.5 Synechocystis sp.......................................................................................................................................... 62 AquaFUELs‐ Taxonomy, Biology and Biotechnology
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8 EUKARYOTIC MICROALGAE........................................................................................................................ 64 8.1 CHLOROPHYTA ................................................................................................................................................ 64 8.1.1 Ostreococcus sp. ......................................................................................................................................... 64 8.1.2 Tetraselmis sp ............................................................................................................................................. 66 8.1.3 Botryococcus braunii.................................................................................................................................. 70 8.1.4 Chlamydomonas reinhardtii ....................................................................................................................... 72 8.1.5 Haematococcus pluvialis ............................................................................................................................ 76 8.1.6 Dunaliella sp............................................................................................................................................... 79 8.1.7 Chlorococcum sp. ....................................................................................................................................... 83 8.1.8 Neochloris oleoabundans............................................................................................................................ 86 8.1.9 Scenedesmus sp........................................................................................................................................... 92 8.1.10 Desmodesmus sp. ........................................................................................................................................ 97 8.1.11 Chlorella sp. ............................................................................................................................................... 99 8.1.12 Parietochloris incisa................................................................................................................................. 109 8.1.13 Prototheca sp. ........................................................................................................................................... 111 8.2 RHODOPHYTA ................................................................................................................................................ 113 8.2.1 Porphyridium cruentum ............................................................................................................................ 113 8.3 BACILLARIOPHYTA ........................................................................................................................................ 115 8.3.1 Benthic diatoms (Amphora; Amphiprora; Cylindrotheca; Navicula; Nitzschia) ...................................... 115 Amphora sp. ..........................................................................................................................................................................115 Amphiprora hyalina ..............................................................................................................................................................116 Cylindrotheca sp. ..................................................................................................................................................................116 Navicula sp. ..........................................................................................................................................................................117 Nitzschia dissipata ................................................................................................................................................................118
8.3.2 Phaeodactylum tricornutum...................................................................................................................... 126 8.3.3 Chaetoceros muelleri................................................................................................................................ 126 8.3.4 Cyclotella cryptica.................................................................................................................................... 136 8.3.5 Odontella aurita........................................................................................................................................ 139 8.3.6 Skeletonema sp.......................................................................................................................................... 141 8.3.7 Thalassiosira pseudonana ........................................................................................................................ 143 8.4 EUSTIGMATOPHYCEAE (PHYLUM HETEROKONTOPHYTA) .............................................................................. 146 8.4.1 Monodus subterraneus.............................................................................................................................. 146 8.4.2 Nannochloropsis sp................................................................................................................................... 148 8.5 HAPTOPHYTA ................................................................................................................................................. 153 8.5.1 Isochrysis sp.............................................................................................................................................. 153 8.5.2 Pavlova sp................................................................................................................................................. 156 8.6 DINOPHYTA ................................................................................................................................................... 159 8.6.1 Crypthecodinium cohnii............................................................................................................................ 159 8.7 LABYRINTHULOMYCETES .............................................................................................................................. 162 8.7.1 Schizochytrium sp. .................................................................................................................................... 162 8.7.2 Thraustochytrium sp. ................................................................................................................................ 164 8.7.3 Ulkenia sp. ................................................................................................................................................ 166 9 MACROALGAE............................................................................................................................................... 167 9.1 CHLOROPHYTA .............................................................................................................................................. 167 9.1.1 Caulerpa sp............................................................................................................................................... 167 Caulerpa racemosa ...............................................................................................................................................................167 Caulerpa taxifolia .................................................................................................................................................................168
9.1.2
Ulva sp. ..................................................................................................................................................... 170
Ulva lactuca..........................................................................................................................................................................170 Ulva rigida............................................................................................................................................................................170
9.1.3 9.1.4
Cladophora sp. ......................................................................................................................................... 178 Codium sp. ................................................................................................................................................ 180
Codium fragile ......................................................................................................................................................................180 Codium parvulum..................................................................................................................................................................180
9.2 RHODOPHYTA ................................................................................................................................................ 183 9.2.1 Chondrus crispus ...................................................................................................................................... 183 9.2.2 Mastocarpus stellatus ............................................................................................................................... 185 9.2.3 Grateloupia turuturu................................................................................................................................. 187 9.2.4 Palmaria palmata ..................................................................................................................................... 189 9.2.5 Solieria chordalis...................................................................................................................................... 191 9.3 PHAEOPHYCEAE (PHYLUM HETEROKONTOPHYTA) ........................................................................................ 193 9.3.1 Alaria esculenta ........................................................................................................................................ 193 9.3.2 Undaria pinnatifida .................................................................................................................................. 195 AquaFUELs‐ Taxonomy, Biology and Biotechnology
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9.3.3 9.3.4
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Ascophyllum nodosum .............................................................................................................................. 197 Fucus sp. ................................................................................................................................................... 199
Fucus serratus.......................................................................................................................................................................199 Fucus spiralis........................................................................................................................................................................199 Fucus vesiculosus..................................................................................................................................................................200
9.3.5 9.3.6
Himanthalia elongata ............................................................................................................................... 202 Cystoseira sp............................................................................................................................................. 204
Cystoseira baccata................................................................................................................................................................204 Cystoseira tamariscifolia ......................................................................................................................................................205
9.3.7 9.3.8 9.3.9
Halidrys siliquosa ..................................................................................................................................... 207 Sargassum muticum .................................................................................................................................. 208 Laminaria, Saccharina, Saccorhiza.......................................................................................................... 210
Laminaria sp. ........................................................................................................................................................................210 Laminaria digitata ...........................................................................................................................................................210 Laminaria hyperborea......................................................................................................................................................211 Laminaria ochroleuca......................................................................................................................................................212 Saccharina latissima .............................................................................................................................................................212 Saccorhiza polyschides .........................................................................................................................................................213
10 OTHER AQUATIC BIOMASS ........................................................................................................................ 221 10.1 EGERIA DENSA ................................................................................................................................................ 221 10.2 EICHHORNIA CRASSIPES................................................................................................................................... 223 10.3 ELODEA CANADENSIS ...................................................................................................................................... 226 10.4 LAGAROSIPHON MAJOR .................................................................................................................................... 228 10.5 LEMNA MINOR ................................................................................................................................................. 231 11 CONCLUDING REMARKS............................................................................................................................. 236 ANNEX I ................................................................................................................................................................... 237
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1 Introduction Algae are a group of organisms that have been generally described as photoautotrophic unicellular or multicellular, mainly water dwelling organisms lacking complex morphological organization. Historically, the prokaryotic blue‐green algae, or cyanobacteria (Class Cyanophyceae), are often included in discussing microalgae, and indeed some cyanobacterial species (Arthrospira or spirulina) hold a prominent position in the biotechnological exploitation of microalgae. There are several main groups of microalgae differing in biochemical constituents, ultrastructure, and life cycle. Some of the characteristics traditionally used for algae classification are the nature of their photosynthetic pigments, storage products, cell wall, presence or absence of flagella and the number of membranes surrounding the chloroplast. More recently classification has been based on comparisons of specific DNA sequences, leading to major revisions in classification of many groups of alga. Recent molecular genetic studies confirmed that photoautotrophic eukaryotes belong to several highly diverse groups of organisms and are the result of different and independent events of secondary endosymbioses. As a consequence algae belong to genetically widely diverting groups of organisms often closer related to nonphotosynthetic organisms than to more distant algal clades (Fig. 1). This fact requires due attention when developing tools such as transformation or genetic engineering etc for microalgae. The most recent results on algal taxonomy, are summarized in detail by the Tree of Life project (http://tolweb.org/tree/) and AlgaeBase (http://www.algaebase.org/), both providing up to date taxonomic information concerning classification of algal species, that is continuously being updated and revised in light of newest results obtained by molecular genetic approaches such as DNA sequence comparisons.
Figure 1 Phylogenetic tree of the eukaryotic organisms (modified from Tree of Life Project
http://tolweb.org/tree/).
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Microalgae reproduction occurs primarily by vegetative (asexual) cell division, although sexual reproduction can occur in many species under appropriate growth conditions.
1.1 Importance of algae and aquatic biomass for biofuels 1.1.1
Suitability of algae as biomass producers
Microalgae are considered fast‐growing photosynthetic organisms and have been reported to reach short term maximal summer productivities of 50 ‐ 60 g per m2 per day in CO2 enriched raceway ponds in Hawaii and California. This corresponds to transformation of 5‐6% of incoming light energy into biomass. Such numbers, as well as productivity data from lab scale experiments have promoted the reputation of microalgae as prime candidates for providing unlimited amounts of cheap biomass as food, fodder or energy. Furthermore many algal strains can produce large amounts of oil or lipid like storage products that can easily be converted into biodiesel (Sheehan et al., 1998). This has been used by some to combine maximal biomass productivity with maximal oil content, yielding phantastic oil productivity numbers that have been exploited for funding intensive research on algal biofuels production. Potential oil productivities of over 100 tons/ha per year were initially predicted. However, none of the large scale long term experiments ever reached the high productivity projections. Current productivity obtained in large scale operations range from 40 – 60 tons of algal biomass production per ha and year, with conservative projections anticipating up to 100 tons of biomass, or 30 tons of biodiesel per ha and year in subtropical or tropical, sunny climates (Scott et al., 2010). Five groups of microalgae were classified as high priority for biofuel production by the US microalgal species program ASP (Sheehan et al., 1998): diatoms (Class Bacillariophyceae), green algae (Class Chlorophyceae), golden‐brown algae (Class Chrysophyceae), prymnesiophytes or haptophytes (Class Prymnesiophyceae), and eustigmatophytes (Class Eustigmatophyceae). However, different classes of macroalgae, as well as further yet less studied microalgal groups may turn out to be equally relevant for successful biomass production from algae. Other aquatic biomass such as water lentils (Lemna), water hyacinth, Elodea and others have also been considered for potential biofuel production due to their significant productivity and their usefulness in treating polluted nutrient rich water bodies. 1.1.2
Sustainability, the strategic advantage of algal biofuels
Land Use ‐ Current biofuels such as oil from soy bean, palm, and rape seed, or ethanol from corn or wheat, suffered from serious setbacks revealed by recent analysis showing their adverse ecological impact and low greenhouse gas reduction potential. A recent statement by UNEP Director, A. Steiner, reads: "biofuels from palm oil grown by Indonesia might never be deemed to be sustainable", due to ongoing destruction of tropical forests for expanding palm oil production. In addition, with yields of less than 500‐5,000 L of biodiesel per hectare (Johnston et al., 2009), those crops require enormous areas of scarce arable land, water and fertilizer and are generally highly work intensive. Life cycle assessments (LCA) indicate that no, or very low, reductions in greenhouse gas emissions can be achieved using such biofuels (Zah et al., 2007), and if they are being produced following conversion of natural ecosystems their GHG emissions surpass those of fossil fuels for years to come (Fargione et al., 2008). The major impact, land use, is often not adequately considered if the strategic implications of expanding biofuels production are taken into account (Searchinger et al., 2008):
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Supplying the world's 2030 liquid fuel demand from dedicated biofuel crops would consume all or most of the available land in the appropriate climatic zones including most remaining natural ecosystems even if the most productive biofuels crops such as palm oil, Miscanthus cellulosic ethanol or sugar cane ethanol or an optimized mix of those crops were planted. Even if a doubling of yields is achieved in the next 20 years, around half of the worlds remaining intact ecosystems will have to be sacrificed to cover the projected liquid fuel demand. Only algae can provide sufficient liquid fuels on a few percent of available dryland areas. Algal biomass if managed properly, may be produced on unproductive desert land, under utilization of ocean or waste water by exploiting and recycling of waste nutrients from municipal and agricultural sources, since no health concerns need to be considered for biofuel production. Algal biofuel production is thus complementary to ongoing efforts to grow lignocellulosic biomass in areas with good soil and water resources, because the microalgae are projected to be grown in those areas where the lignocellulosic or oil seeds crops will not perform well (Brown, M. Lewis, “Biodiesel from Microalgae: Complementarity in a Fuel Development Strategy”, NREL, http://www.nrel.gov/docs/legosti/old/5715.pdf ). Algae may produce sufficient biofuels on less than 10% of available drylands even under current productivity estimates, often using the most unproductive areas like salt flats or degraded dryland soils. They may also deliver additional environmental services such as wastewater treatment or exhaust gas detoxification. Table 1 ‐ Comparison of land use impact of various biofuel crops. *For all crops it is assumed that 50% of biomass energy is used for process energy and all nutrients will be recycled to the maximum possible, according to the state of the art of sugar cane ethanol production. In Miscanthus this 50% are removed from the claimed ethanol yield, since no leftover biomass is available for process energy. The crops marked in italics, as well as algae, are experimental as no actual production in the large scale has been demonstrated. (Land areas are derived from Ito and Oikawa, 2004; biofuel productivities are derived from Johnston et al., 2009).
Land type
Area (mio km2)
Tropical and subtropical evergreen forest Tropical and subtropical dry forest Tropical Savanna, Woodland Mid lattitude forests, abandoned croplands
Warm Shrubland/grassland or desert
Natural Productivity (tons of carbon fixed per
Crop and biofuel yield
hectare and year)
(tons per ha / GJ per ha)
Palmoil (5 / 189) Jatropha‐ oil (1.5 / 56.7 )
% area of corresponding ecosystem required to cover 2030 demand
10.5
10.7
110%!!!
4.7
7.67
6.7
6.65
Cane‐ethanol (4.34 / 116)
270 %
14
5.30
Miscanthus cellulosic ethanol* (4.4 / 120)
95 %
33
1 – 3.50
Algae‐ oil (20 / 756)
5.4 – 8.2 %
765%!!
Other Environmental Services ‐ Interestingly, even under desert conditions the water footprint of algal biomass production is lower than that of irrigated maize ethanol production, calculated on the water input per unit of energy created. Our estimates suggest that it requires less than 0.5 million liters of water to produce one 1 ton of dry algal biomass, and this may be waste‐ or seawater. It can take about 3 million AquaFUELs‐ Taxonomy, Biology and Biotechnology
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liters of water to produce 1 ton of rice and about 2 million liters to produce 1 ton of soybeans (www.clw.csiro.au/issues/water/water_for_food.html and www.gdrc.org/uem/footprints/water‐ footprint.html). However, significant investments into (waste)‐water and CO2 infrastructure would be required to achieve the necessary global algae biomass production potential. Properly planned algal biomass production facilities may recover and reuse most of the nutrients applied, minimizing eutrophication impact, and this in contrast to intensive agriculture where nutrient runoff and escaping nitric oxide pose serious problems. In fact, algae have been shown to be able to treat successfully any kind, even the most problematic, forms of wastewater. Pesticide use in algal cultivation is expected to be minimal. Even a small proportion of the algal biomass required for energy purposes would provide sufficient protein to replace all the soybean cultivation capacity installed for feed production, resulting in reduced deforestation or 'negative indirect land use changes' (Searchinger et al., 2008). While algal biomass production may never achieve the low production costs of other agricultural commodities, full accounting of above and additional environmental services may result in a balance favouring the algal fuels. This point requires intensive investigation progressing far beyond currently used LCA models (Stephenson et al., 2010), starting with defining those production parameters and system boundaries that will actually deliver the above mentioned environmental advantages.
1.2 Rationale of the document This document summarizes in short current views and prospects on the potential contribution of algae to biofuel production. Recent predictions and calculations, both at the EU level and in the US (A USDA Regional Roadmap to Meeting the Biofuels Goals of the Renewable Fuels Standard by 2020, USDA Strategic Biofuels Production report, June 2010), do not incorporate a significant algae potential into their projections for the next 10 years. This is dictated by the fact that algae experts and external observers disagree about the true potential of algal biofuels production relating to economic and environmental sustainability, and any given time frame for achieving competitive algal biofuel production is speculative at the best. Given the high complexity of algal taxonomy and evolutionary relationships, this document was conceived as an instrument to place the algae that have arisen an interest for biofuel production within the correct frame. The list of algae proposed is based on the literature concerning biofuel production, on the commercially produced algae and on the feedback from the questionnaire in deliverable 1.1. Detailed description of biotechnology is provided only for pivotal taxa or group of taxa, that are actually produced at least at pilot scale. These taxa represent the reference model for those taxa that are not currently exploited. It was beyond the scope of this document to propose any kind of new or revised taxonomy of algae. The classification reported is based on AlgaeBase (http://www.algaebase.org/) and Tree of Life project (http://tolweb.org/tree/). The other aquatic biomasses species reported are all invasive weeds that have been proposed as biofuel crops. The classification reported is based on ITIS Catalogue of Life 2010 (http://www.catalogueoflife.org/annual‐checklist/2010/search/all) and US Department of Agriculture PLANTS Database (http://plants.usda.gov/).
1.3 Target groups Since the major gaps in knowledge concerning algal biofuels are the lack of operating pilot scale and commercial algal biofuels production facilities, the target audience for this report are all interested AquaFUELs‐ Taxonomy, Biology and Biotechnology
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stakeholders in Europe’s energy, agricultural and environmental policy, such as policy makers, NGOs, research infrastructures, interested industries and financial institutions. The conclusion of this report should be an appeal for rapid and significant investments into algae related biomass production in the form of public or privately managed pilot or production facilities with the aim of testing a multitude of operational parameters for increasing yields and sustainability while reducing production costs of algal biomass as much as possible. Taking into account the long term multi trillion Euro per year energy and by‐ product market, this report aims at convincing all involved potential stakeholders and investors to designing long term strategies for funding an algae based fuel and biomass development program that may provide the necessary insight on their undoubtedly huge potential.
1.4 Problems incurred It is a disturbing fact that today no results on true sustained biomass production yields for biofuels production are available, and no functional biofuel production plant is accessible anywhere around the world. Thus all recent publications in the field, be it yields, economics or LCA, remain pure speculation and demand greatest care in their interpretation. During the last 10 years algal biofuels companies, driven by large investments from venture capital, have aimed to demonstrate a potential for rapidly achieving economic profitability. This trend lead to high degrees of secrecy surrounding mysterious processes developed whose technical and scientific soundness cannot be confirmed nor discarded due to lacking access to raw data and facilities. The summary presented below therefore relies on publications and patents released by mostly academic institutions and a few open minded companies feeling that little in terms of technology and biology in the process of algal biomass production deserves this degree of secrecy. Nevertheless it cannot be excluded that certain secret breakthroughs may have been achieved recently that would put the state of the art significantly ahead of what is being presented here.
1.5 Common erroneous "myth" In contrast to often voiced opinions, algae are not significantly more efficient in biomass production than other plants grown under optimal conditions. The most common error is comparing biomass doubling time or specific growth rates, which indicate the rate of biomass accumulation under exponential growth conditions, where indeed algae and cyanobacteria may multiply several times per day. However, those conditions are possible under very low biomass densities only that are not applicable to large scale algal cultivation since actual biomass produced per day is in fact lower than in cultures with higher biomass densities, where all the incoming light is captured by algae and used for photosynthesic biomass production. If such doubling times of exponentially growing cultures are being applied to denser cultures (which could be done with heterotrophic organisms by increasing the food input) in fact very easily fantastic daily growth rates can be assumed. However other than in heterotrophic culture conditions the one and only energy source for growth of algae is incoming light energy that is transformed with an efficiency of around 3% into biomass. Under absolutely optimized conditions in terms of temperature, light intensity, mixing and CO2 supply, higher photosynthetic efficiencies of up to 7% may be achieved, however under exponential increase in bioreactor and maintenance costs that are generally claimed not to be covered by increased yield, and also require far higher energy inputs leading to a negative ratio between energy input and gain in form of algal biomass for example in tubular photobioreactors (Jorquera et al. 2010).
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2 Criteria for strain selection The objective was to agree on criteria for both micro‐ and macro algae on which species can be selected for their suitability for biofuel production. These criteria should ideally be quantitatively measurable. For all criteria one should keep an outdoor, large scale system in mind, because for lab (scale) experimentations different criteria may apply.
2.1 Productivity This includes productivity of biomass and of specific biomass components (e.g. lipids). In order to be able to compare algae with traditional crops and with each other in terms of productivity, the most objective criterion is to use photosynthetic efficiency as a measure for productivity. Basically this means the % of available light (energy) that is converted into biomass or specific biomass components. In this way all species and all cultivation systems can be compared.
2.2 Robustness This is a rather vague term which includes resistance to many extreme conditions. This criterion can be best assessed using table in which the resistance to these several conditions is scored. Table 2‐ Conditions of robustness.
Condition pH Oxygen concentration Temperature
Salinity
Organic contaminants
Relevant for Reduce risk of infection CO2 transfer Closed photobioreactors Outdoor cultivation Open water cultivation
Range i.e. 10
>20% Large range to accommodate day/night and seasonal fluctuation (e.g. 10 – 40 °C) Cultivation in fresh / sea / e.g 0‐10% salinity brackish water Reduce risk of infection Ability to grow on Concentration of organic wastewater / flue gas contaminants that still allows good growth
2.3 Harvestability For microalgae this will mainly be the sedimentation rate and the possibilities for induced‐ or auto flocculation. For macroalgae this includes the possibilities for mechanical harvesting or harvesting by hand.
2.4 Biomass composition This should include a breakdown of the total biomass composition in: • total caloric value of the biomass (for burning it), • % lipids and lipid composition (for biodiesel), AquaFUELs‐ Taxonomy, Biology and Biotechnology
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• • •
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% starch and carbohydrate composition (for bio ethanol and to identify higher value byproducts (i.e. agar), % protein and protein composition (soluble/insoluble for food/feed purposes), presence of heavy metals or toxins (specification).
2.5 Processability / extractability This should include relevant aspects for biorefinery, such as the cell volume, thickness/toughness of the cell wall and the presence of tough fibers (macroalgae) and the moisture content. A measure for this could be the energy input per gram of dry weight necessary for full biorefinery.
2.6 Added value of coproducts Does the organism produce any by‐ or co‐product that have an intrinsic added value, such as carotenoids. This is important to reduce the costs of the final biofuel product. Here a specification of the compounds and their expected added value per gram of dry biomass should be indicated.
2.7 Local origin of strains The use of locally selected strains may be of significance both for ease of management and for reasons of sustainability Based on criteria of the 'Roundtable on Sustainable Biofuels' (http://rsb.epfl.ch/). Non‐native potentially invasive biofuels crops should not be used in open cultivation systems, and adherence to this rule will require the identification and use of locally isolated algal strains. Furthermore such strains may have unique adaptations to the local climate, water and possible parasites that imported or even laboratory grown strains may not have.
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3 Biology of algae Algae are an assemblage of organisms that have been generally described as photoautotrophic unicellular or multicellular organisms lacking complex morphological organization, and as such have been reclassified several times in recent biology according to technical advances on the basis of differences in subcellular organization, or later of molecular genetic characteristics that will allow, when sufficient data will be available, precise determination of evolutionary relationships. Historically, algae included also cyanobcateria, prokaryotic oxygenic phototrophs. Many microalgae grow quite rapidly and their reproduction occurs primarily by vegetative (asexual) cell division, although sexual reproduction can occur in many species under appropriate growth conditions. There are several main groups of microalgae, which differ primarily in pigment composition, biochemical constituents, ultrastructure, and life cycle. Five groups were of primary importance: diatoms (Bacillariophyta), green algae (Chlorophyta), Prymnesiophyta or Haptophyta and Eustigmatophyta together with the prokaryotic blue‐green algae, or cyanobacteria. Recent molecular genetic studies confirmed that photoautotrophic eukaryotes belong to several highly diverse groups of organisms and are the result of different and independent events of secondary endosymbioses. As a consequence algae are a genetically widely diverting group of organisms, a fact that will require due attention when developing tools such as transformation or genetic engineering etc.
3.1 Cyanobacteria Cyanobacteria are prokaryotic photoautotrophic microorganisms that can be found in almost every environment, from oceans to freshwater to bare rock to soil. Though the prokaryotic Cyanobacteria (commonly referred to as blue‐green algae) were traditionally included as "algae" in older textbooks, many modern sources regard this as outdated as they are now considered to be bacteria. Classification ‐ The cyanobacteria were traditionally classified by morphology according to the International Code of Botanical Nomenclature into five orders: Chroococcales, Pleurocapsales, Oscillatoriales, Nostocales and Stigonematales. Starting from the 1970s cyanobacteria have been classified also according to the International Code of Nomenclature of Bacteria and they were treated in the Bergey’s Manual of Systematic Bacteriology Volume 3 of the 1989 edition, then updated in volume I of the edition of 2004, where cyanobacterai are subdivided in five subsections, I‐V, corresponding to the orders of the Botanical Code, except that the Prochlorales have been included within the cyanobacteria and precisely in the I subsection (Komárek and Anagnostidis, 1986; Anagnostidis and Komárek, 1988; Castenholz and Waterbury, 1989; Komárek and Anagnostidis, 1989; Anagnostidis and Komárek, 1990; Castenholz, 2001;Oren, 2004; Wilmotte and Herdman, 2001; Herrero and Flores (eds), 2008). Cell structure ‐ The first two subsections include unicellular cyanobacteria (Castenholz and Waterbury, 1989; Castenholz, 2001). The members of Chroococcales are unicellular cyanobacteria that reproduce by binary fission or budding. Cells are coccoid or rod shaped and can vary in length from 0.5 to 30 μm. Division can occur in one to three successive planes, so that cells can be single or in colonies. The classic taxonomic criterion has been the cell morphology and the plane of cell division. In AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Pleurocapsales, cyanobacteria reproduce by multiple fission which generates small spherical cells named baeocytes, that can be motile or not according to the genus (Herdman and Rippka, 1988b). In unicellular forms there is only multiple fission, whereas in colonial forms after binary fission on different planes some of the cells undergo multiple fission. The remaining sections include filamentous cyanobacteria. In Oscillatoriales (Castenholz and Waterbury, 1989; Castenholz, 2001), the cells are uniseriately arranged and do not form specialized cells (akinetes and heterocysts). They reproduce by binary fission in a single plane. Filament diameter varies from 0.4 to 100 μm. Outside the cell wall a sheath may be present. In this case terminal hormogonia of short length can glide out of the sheath and eventually form new sheaths. New filaments (or trichomes) are originated from fragmentation in correspondence to a dead cell or certain cells (necridial cells) are purposely destined to die. In Nostocales and Stigonematales (Castenholz and Waterbury, 1989; Castenholz, 2001) the cells have the ability to differentiate cells like heterocysts and akinetes. Nostocales are filamentous cyanobacteria dividing only by binary fission in one plane, though some genera produce false branching. Filaments may be composed of cells of uniform diameter or by cells with decreasing diameter towards the end of the filament (tapering trichomes). Heterocysts may be terminal and intercalary or only terminal in different genera. Motile trichomes (hormogonia) can be formed for dispersion in some genera (Herdman and Rippka, 1988b). Resistance cell (akinetes) may be formed under unfavorable conditions (Herdman and Rippka, 1988a). Stigonematales, unlike Nostocales, includes species with truly branched trichomes. Within this class there is the maximum degree of complexity and differentiation of all the cyanobacterial groups. Longitudinal or oblique cell division occurs in addition to tranverse division, so that periodic true branching and, in some cases, multiseriate trichomes are formed. Hormogonia may be formed, even if reproduction occurs mainly by random breakage of the trichome. Akinets can be produced in some genera. Heterocysts are both intercalary and terminal. Cell diameter varies within a “trichome” as secondary branches are usually narrower. In cyanobacteria (Castenholz and Waterbury, 1989), generation times are usually higher than 24 h, though some unicellular and oscillatorian strains can duplicate in 4 h. Some genera can have complex morphogenetic cycles including filamentation, aseriate phases, dispersal through hormogonia and akinetes production. Cyanobacteria share the basic cell characteristics with the other Bacteria (Stanier and Cohen‐ Bazire, 1977; Castenholz and Waterbury, 1989). The cell wall is of the Gram‐negative type, though the peptidoglycan layer is considerably thicker than in the other Gram‐negative bacteria. Many cyanobacteria have a sheath or glycocalix or capsule or gel, mucilage or slime outside the outer membrane of the cell wall, mainly composed of polysaccharides. Some sheaths can have a microfibrillar structure and can become laminated with aging of the trichome. Some cyanobacteria (and many hormogonia) can contain gas‐vesicles that allow buoyancy in the water column. Cyanobacteria have an elaborate and highly organized system of internal membranes which function in photosynthesis (thylakoids) (Stanier and Cohen‐Bazire, 1977; Castenholz and Waterbury, 1989). The lipophilic pigments chlorophyll a (both reaction centers and antenna) and photosynthetic carotenoids are located within the thylakoids, while the hydrophilic antenna pigments (allophycocyanin‐APC‐, phycocyanin ‐PC‐ and, where they are present, phycoerythrin –PE‐ or phycoerythrocyanin ‐PEC) are located in the phycobilisomes which are attached to the outside of the thylakoid membranes. The phycobilisome is haemidiscoidal and is composed of stacks of biliproteins in AquaFUELs‐ Taxonomy, Biology and Biotechnology
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the order (from inside to outside) APC, PC, PE or PEC. Genera belonging to the former group of the Prochlorales lack phycobilisomes and have chlorophyll b as antenna pigment. The cyanobacterium Acaryochloris marina has been reported to contain chlorophyll d instead of chlorophyll a as light harvesting pigment, so that its photosynthetic process depends on far‐red light (710‐718 nm) (Miyashita et al., 2003). The reserve carbohydrate is glycogen (Stanier and Cohen‐Bazire, 1977; Castenholz and Waterbury, 1989). Cyanobacteria contain also cyanophycin, a nitrogen reserve polymer made of arginine and aspartic acid, polyphosphate granules and carboxisomes, that are a cell reserve of the photosynthesis key enzyme rubisco (ribulose‐1,5‐biphosphate carboxylase). Some cyanobacteria also contain poly‐β‐ hydroxybutyrate granules. Physiology ‐ Photosynthesis in cyanobacteria (Wolk, 1973; Stanier and Cohen‐Bazire, 1977; Castenholz and Waterbury, 1989)uses water as an electron donor and produces oxygen as a by‐product. This water‐oxidizing process is accomplished by coupling the activity of photosystem (PS) II and I (Z‐ scheme). Under anaerobic conditions some genera (belonging to the I and III subsections) are able to carry out anoxygenic photosynthesis using only PS I to carry out cyclic photophosphorylation and obtain ATP, and using electron donors other than water (hydrogen sulfide or thiosulphate) (Cohen et al., 1975; Garlick et al., 1977). Carbon dioxide is reduced to form carbohydrates via the Calvin cycle. During respiration reduced NADP is obtained through the pentose phosphate cycle. The plasma membrane contains only components of the respiratory chain, while the thylakoid membrane hosts both respiratory and photosynthetic electron transport. Most cyanobacteria are obligate photoautotrophs, but some species can grow as heterotrophs in the dark at the expense of glucose, fructose or sucrose. Under anaerobic conditions, some species can perform lactate fermentation (Oren and Shilo, 1979). Nitrogen fixation occurs both in heterocystous cyanobacteria and in some non‐hetrocystous cyanobacteria. To avoid contact of nitrogenase with oxygen (and then its permanent inactivation) these latter cyanobacteria adopt a temporal separation between the photosynthetic and the nitrogen fixation processes (Bergman et al., 1997). Increased respiration rates allow to control the oxygen concentration inside the cell, due to diffusion, necessary to carry out cell metabolism. In heterocystous forms, the nitrogen fixation process is spatially separated from the oxygenc photosynthesis. Nitrogen fixation is carried out in specialized cells, the heterocysts (Adams and Duggan, 1999). These have many characteristics that allow to reduce diffusion of oxygen, such as a thick cell wall surrounded by a complex external envelope and a reorganization of the photosynthetic apparatus: lack of PS II to avoid internal oxygen production, presence of PS I to obtain ATP through cyclic photophosphorilation. Reducing power is obtained from vegetative cells in the form of sugars. Molecular nitrogen is fixed into ammonia and immediately converted to organic form, usually as glutamine. As nitrogen fixation is a very energy‐consuming process, nitrogenese is produced and heterocysts are differentiated only in the absence of combined nitrogen in the environement surrounding the cell. Ecology ‐ Cyanobacteria are the only group of organisms that are able to reduce nitrogen and carbon in aerobic conditions, a fact that may be responsible for their evolutionary and ecological success (Whitton and Potts (eds), 2000). They contribute significantly to global ecology and the oxygen cycle. The large amounts of oxygen in the atmosphere originally derive from the activities of ancient cyanobacteria. The tiny marine cyanobacterium Prochlorococcus was discovered in 1986 (Chisholm et AquaFUELs‐ Taxonomy, Biology and Biotechnology
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al., 1988) and, together with the picoplanktonic cyanobacteria, accounts for up to half of the primary production of waters, from oligotrophic open ocean to estuarine ecosystems. Due to their ability to fix nitrogen in aerobic conditions they are often found as symbionts (Rai et al. (eds), 2002) with a number of other groups of organisms such as fungi (lichens), corals, pteridophytes (Azolla), angiosperms (Gunnera), protists (including some diatoms), and sponges. The rice paddies of Asia rely on nitrogen‐fixing cyanobacteria as fertilizers, both free biomass and symbiont to the fern Azolla. They can occur as planktonic cells (thanks to the buoyancy ability) or form phototrophic biofilms in freshwater and marine environments, they occur in damp soil, or even temporarily moistened rocks in deserts (Whitton and Potts (eds), 2000). Some live in the fur of sloths, providing a form of camouflage. Aquatic cyanobacteria are probably best known for the extensive and highly visible blooms that can form in both freshwater and marine environments. The association of toxicity with such blooms has frequently led to the closure of recreational waters when blooms are observed. Certain cyanobacteria produce cyanotoxins (Chorus and Bartram, 1999) including neurotoxins, hepatotoxins, cytotoxins, and endotoxins. Examples of cyanotoxins are anatoxin‐a, anatoxin‐as, aplysiatoxin, saxitoxin, cylindrospermopsin, microcystins, nodularin. These toxins can be dangerous to humans and animals. Several cases of human poisoning have been documented. Recent studies suggest that significant exposure to high levels of BMAA a non‐proteic aminoacid produced by many cyanobacteria could be among the causes of neurodegenerative diseases such as Amyotrophic Lateral Sclerosis. Genome sequencing ‐ The unicellular cyanobacterium Synechocystis sp. PCC6803 was the third prokaryote and first photosynthetic organism whose genome was completely sequenced (Kaneko et al. 1996) . It continues to be an important model organism. Today over 40 complete cyanobacterial genomes are known (www.ncbi.nlm.nih.gov). The smallest genomes have been found in Prochlorococcus spp. (1.7 Mb) and the largest in Nostoc punctiforme (9 Mb). Those of Calothrix spp. are estimated at 12‐15 Mb, as large as yeast. Relationship to chloroplasts ‐ Chloroplasts found in eukaryotes (algae and plants) likely evolved from an endosymbiotic relation with cyanobacteria. This endosymbiotic theory is supported by various structural and genetic similarities. Primary chloroplasts are found among the "true plants" or green plants as well as among the red algae and glaucophytes, marine species which contain phycobilins. It now appears that these chloroplasts probably had a single origin, in an ancestor of the clade called Primoplantae. Other algae likely took their chloroplasts from these forms by secondary endosymbiosis or ingestion.
3.2 Chlorophyta (Green Algae) The green algae are a large group of algae from which the embryophytes (higher plants) emerged (Jeffrey et al., 2004). The group including both green algae and embryophytes is monophyletic (and often just known as kingdom Plantae). The green algae include unicellular and colonial flagellates, usually but not always with two flagella per cell, as well as various colonial, coccoid, and filamentous forms. In the Charales, the closest relatives of higher plants, full differentiation of tissues occurs (Thomas, 2002). There are about 6,000 species of green algae. Many species live most of their lives as single cells, while other species form colonies or long filaments. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Some species of green algae, particularly of genera Trebouxia and Pseudotrebouxia (Trebouxiophyceae), can be found in symbiotic associations with fungi to form lichens. In general the fungal species that partner in lichens can not live on their own, while the algal species is often found living in nature without the fungus. Prasinophyceae are a class of primitive eukaryotic marine green algae (Sym and Pienaar, 1993). Their best known genus is Ostreococcus, which is considered to be the smallest (ca. 0.95 μm) free‐ living eukaryote and which has been detected in marine samples around the world (Courties et al., 1994). Prasinophyceae are thought to have low cellular complexity, that is, they possess single, multiple or no flagella and contain only a single chloroplast and a single mitochondrion. They also have very small genomes for a eukaryote (about 12 Mbp), and the genomes of two Ostreococcus species, taurii and lucimarinus, have been completely sequenced. It has been suggested that a prasinophyceae‐ like flagellate was the ancestor to Chlorophyta and Streptophyta (Kapraun, 2007). A study of photosynthetic gene‐sequence diversity (rbcL) in the Gulf of Mexico indicated that Prasinophyceae are particularly prevalent at the Subsurface Chlorophyll Maximum (SCM) (Warwick et al., 2003) and several different ecotypes of Ostreococcus have been detected in the environment (Guillou et al., 2004). These ecotypes are distinguished by their adaptation to light intensities. The Chlorophyceae are one of the classes of green algae, distinguished mainly on the basis of ultrastructural morphology. For example the chlorophycean CW clade, and chlorophycean DO clade, are defined by the arrangement of their flagella. Members of the CW clade have flagella that are displaced in a "clockwise" (CW, 1–7 o'clock) direction eg. Chlamydomonadales. Members of the DO clade have flagella that are "directly opposed" (DO, 12–6 o'clock) e.g. Sphaeropleales. They share many similarities with the higher plants, including the presence of asymmetrical flagellated cells, the breakdown of the nuclear envelope at mitosis, and the presence of phytochromes, flavonoids, and the chemical precursors to the cuticle (Raven et al., 2005). Cell structure ‐ Almost all forms have chloroplasts. These contain chlorophylls a and b, giving them a bright green colour (as well as the accessory pigments beta carotene and xanthophylls), and have stacked thylakoids (van den Hoek et al., 1995). All green algae have mitochondria with flat cristae. The storage product for members of this group is true starch, amylose, and amylopectin (α‐1,4‐linked polyglucans), and is found inside the chloroplasts. The starch (seen as whitish granules with the TEM) can often be observed surrounding the pyrenoid, a distinct spherical structure embedded in the chloroplast. There may be more than one pyrenoid or the prenoid is not always present (e.g., Ankistrodesmus and Tetraedron) or the pyrenoid is lacking. In most representative taxa, the cells are surrounded by a cellulose cell wall (Wehr and Sheath, 2003). Some taxa may also have chitin or sporopollenin deposited on the wall. This gives added strength and is thought to help prevent desiccation. Some taxa have wall ornamentation, such as scales, a rough texture, thick walls with distinct layers, warts, ridges, and spines. The Volvocales usually have cell walls, loricae, or gelatinous matrices and the main component of the cell walls is glycoprotein, rather than cellulose. The flagellated green microalgae can have from one to eight isokont flagella. The Chlorophyta macroalgae share the following common characteristics: flagella of swimming cells in pairs or multiples of two; stellate structure linking nine pairs of microtubules at basal body transition zone; thylakoids single or stacked; plastid with two membranes without periplastid endoplasmic reticulum; starch inside plastid; glycolate dehydrogenase present; cell wall, when present, of cellulose; cell division without phragmoplast. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Origin ‐ The chloroplasts of green algae are bound by a double membrane, so presumably they were acquired by direct endosymbiosis of cyanobacteria. A number of cyanobacteria show similar pigmentation, but this appears to have arisen more than once, and the chloroplasts of green algae are no longer considered closely related to such forms. Instead, the green algae probably share a common origin with the red algae. Phylogeny ‐ The orders outside the Chlorophyta are often grouped as the division Charophyta, which is paraphyletic to higher plants, together comprising the Streptophyta. Sometimes the Charophyta is restricted to the Charales, and a division Gamophyta is introduced for the Zygnematales and Desmidiales. In older systems the Chlorophyta may be taken to include all the green algae, but taken as above they appear to form a monophyletic group. One of the most basal green algae is the flagellate Mesostigma, although it is not yet clear whether it is sister to all other green algae, or whether it is one of the more basal members of the Streptophyta. Reproduction ‐ Most green algae can proliferate vegetatively by cell division, often the mother cell can divide into up to 16 offspring before releasing them. They often can profliferate sexually whereby haploid algae cells of opposing mating type (containing only one copy of their DNA) can fuse with other haploid cells to form diploid zygotes. They can also follow a reproduction cycle called alternation of generations. Reproduction varies from fusion of identical cells (isogamy) to fertilization of a large non‐motile cell by a smaller motile one (oogamy). However, these traits show some variation, most notably among the basal green algae, called prasinophytes. Some taxa produce motile cells (planospores). Planospores may be asexual zoospores or sexual gametes. Aplanospores (nonmotile cells) may be also produced. When filamentous algae conjugate, they form bridges between cells, and leave empty cell walls behind that can be easily distinguished under the light microscope. The species of Ulva are reproductively isomorphic, the diploid vegetative phase is the site of meiosis and releases haploid zoospores, which germinate and grow producing a haploid phase alternating with the vegetative phase.
3.3 Rhodophyta (Red Algae) The Rhodophyta are a distinct eukaryotic lineage characterized by the accessory photosynthetic pigments phycoerythrin, phycocyanin and allophycocyanin arranged in phycobilisomes, and the absence of flagella and centrioles (Woelkerling, 1990). This is a large assemblage of between 2500 and 6000 species in about 670 largely marine genera (Woelkerling, 1990) that predominate along the coastal and continental shelf areas of tropical, temperate and cold‐water regions (Lüning, 1990). Red algae are ecologically significant as primary producers, providers of structural habitat for other marine organisms, and their important role in the primary establishment and maintenance of coral reefs. Red algae are common and widespread, and ecologically important. Cell structure ‐ Red algae have a number of general characteristics that in combination distinguish them from other eukaryotic groups: • • •
absence of flagella and centrioles, floridean starch as a storage product and the storage of starch in the cytoplasm, phycoerythrin, phycocyanin, and allophycocyanin as accessory pigments,
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unstacked thylakoids in plastids, no chloroplast endoplasmic reticulum.
The rhodophyta exhibit the following common characteristics: they are unicellular to multicellular (up to 1 m), mostly free‐living but in some cases parasitic or symbiotic, with chloroplasts containing phycobilins. Cell walls are made of cellulose with mucopolysaccharides (mainly agars and carrageenans) penetrated in many red algae by pores mostly blocked by proteins (complex referred to as pit connections). Their mitochondria have flat cristae sometimes associated with forming faces of dictyosomes. Thylakoids are single, with phycobilisomes, plastids with peripheral thylakoid. During mitosis, the nuclear envelope mostly remains intact but some microtubules of spindle extend from noncentriolar polar bodies through polar gaps in the nuclear envelope. Phylogeny ‐ Traditionally the red algae were divided into two Classes the Bangiophyceae and Florideophyceae. Alternatively a single Class, the Rhodophyceae and two Subclasses, Bangiophycidae and Florideophycidae are used. Based on ultrastructure and molecular evidence the Bangiophyceae is now accepted as a paraphyletic group, while the Florideophyceae is considered to be monophyletic based on two synapomorphic characters ‐ presence of a filamentous gonimoblast and tetrasporangia (Garbary and Gabrielson, 1990 [and references within], Ragan et al., 1994). Reproduction ‐ They usually have separated phases of vegetative growth and sexual reproduction.
3.4 Heterokontophyta The Heterokontophyta are a major line of eukaryotes. Most are algae, ranging from the giant multicellular kelp to the unicellular forms. The name heterokonts refers to the motile life cycle stage, in which the flagellate cells possess two different shaped flagella (Leipe et al., 1994; Patterson, 1989). Cell structure ‐ Heterokont algae are surrounded by four membranes, which are counted from the outermost to the innermost membrane. The first membrane is continuous with the host's chloroplast endoplasmic reticulum, or cER. The second membrane presents a barrier between the lumen of the endoplasmic reticulum and the primary endosymbiont or chloroplast, which represents the next two membranes, within which the thylakoid membranes are found. This arrangement of membranes suggest that heterokont chloroplasts were obtained from the reduction of a symbiotic red algal eukaryote, which had arisen by evolutionary divergence from the monophyletic primary endosymbiotic ancestor that is thought to have given rise to all eukaryotic photoautotrophs. The chloroplasts usually contain chlorophyll a and chlorophyll c, and usually the accessory pigment fucoxanthin, giving them a golden‐brown or brownish‐green color. Many heterokonts are unicellular flagellates, and most others produce flagellate cells at some point in their life‐cycle, for instance as gametes or zoospores. The name heterokont refers to the characteristic form of these cells, which typically have two unequal flagella. The anterior or tinsel flagellum is covered with lateral bristles or mastigonemes, while the other flagellum is whiplash, smooth and usually shorter, or sometimes reduced to a basal body. The flagella are inserted subapically or laterally, and are usually supported by four microtubule roots in a distinctive pattern. Mastigonemes are manufactured from glycoproteins in the cell's endoplasmic reticulum before being transported to its surface. When the tinsel flagellum moves, these create a backwards current, pulling the cell through the water or bringing in food. The mastigonemes have a peculiar tripartite AquaFUELs‐ Taxonomy, Biology and Biotechnology
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structure, which may be taken as the defining characteristic of the group, thereby including a few protists that do not produce cells with the typical heterokont form. They have been lost in a few lines. Origins ‐ Most basal heterokonts are colorless. This suggests that they diverged before acquisition of chloroplasts within the group. Fucoxanthin‐containing chloroplasts are also found among the haptophyta. These two groups may have a common ancestry, and possibly also a common phylogenetic history with cryptophyta. This may be interpreted as suggesting that the ancestral heterokont was an alga, and all colorless groups arose through loss of the secondary endosymbiont and its chloroplast. Phylogeny ‐ As noted above, classification varies considerably. Originally the heterokont algae were treated as two divisions, first within the kingdom Plantae and later the Protista. In this scheme, however, the Chrysophyceae are paraphyletic to both other groups. As a result, various members have been given their own classes and often divisions. Recent systems often treat these as classes within a single division, called the Heterokontophyta, Chromophyta or Ochrophyta. This is not universal, however ‐ for instance Round et al. (1990) treat the diatoms as a division. The discovery that oomycetes and hyphochytrids are related to these algae, rather than fungi as previously thought, has led many authors to include them among the heterokonts. Should it turn out that they evolved from colored ancestors, the group would be paraphyletic in their absence. Once again, however, usage varies. Patterson (1999) named this extended group the stramenopiles, characterized by the presence of tripartite mastigonemes, mitochondria with tubular cristae, and open mitosis. He used the stramenopiles as a prototype for a classification without Linnaean ranks. Their composition has been essentially stable, but their use within ranked systems varies. Cavalier‐Smith (1981) treats the heterokonts as identical in composition with the stramenopiles; this is the definition followed here. He has proposed placing them in a separate kingdom Chromalveolata, together with the haptophytes, cryptomonads and alveolates. This is one of the most common revisions to the five‐ kingdom system, but has not been generally adopted, partly because some biologists doubt their monophyly. A few treat the Chromalveolata as identical in composition with the heterokonts, or list them as a kingdom Stramenopila. 3.4.1
Phaeophyceae (Brown algae)
The class of the Phaeophyceae (Guiry and Guiry, 2007), or brown algae, is a large group of mostly marine multicellular algae, including many seaweeds of colder Northern Hemisphere waters. Brown algae are unique among heterokonts in developing into multicellular forms with differentiated tissues, but they reproduce by means of flagellate spores and gametes, which closely resemble other heterokont cells. Genetic studies show their closest relatives to be the yellow‐green algae. They play an important role in marine environments both as food, and for the habitats they form. For instance Macrocystis, a member of the Laminariales or kelps, may reach 60 m in length, and forms prominent underwater forests. Another example is Sargassum, which creates unique habitats in the tropical waters of the Sargasso Sea. Many brown algae such as members of the order Fucales are commonly found along rocky seashores. Some members of the class are used as food for humans. Worldwide there are about 1500‐2000 species of brown algae. Some species are of sufficient commercial importance, such as Ascophyllum nodosum, that they have become subjects of extensive research in their own right (Senn, 1987; van den Hoek, 1995). AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Algal structure ‐ Filamentous, syntagmatic or parenchymatous; cell wall present, containing alginate compounds and cellulose; plasmodesmata or pores between cells in parenchymatous forms; chloroplasts with girdle lamella; outer chloroplast endoplasmic reticulum membrane with direct membrane connection to the outer nuclear envelope membrane; plastid DNA with ring‐type genophore; eyespots present or absent; plastid pigments include chlorophylls a and c1 and c2, fucoxanthin, and violaxanthin; swimming cells with two flagella usually inserted laterally, one anteriorly directed, one posteriorlydirected; usually four microtubular kinetosome roots but no striated kinetosome root (rhizoplast); flagellar transitional helix typically with 6 gyres located above the major transitional plate; no paraflagellar rod; little to substantial tissue differentiation occurring in parenchymatous forms.; macroscopic or microscopic, some polysiphonous; some form crusts, cushions or are hollow and others grow to form large leathery fronds (Jones, 1962). Evolutionary history ‐ Phaeophyta evolved from the phaeothamniophyceae between 150 and 200 million years ago. Claims that earlier (Ediacaran) fossils are brown algae have since been dismissed (Loeblich, 1974; Medlin et al., 1997; Lee, 2008). The lineages of brown algae diverged in the following order, from oldest to youngest: Dictyotales; Sphacelariales; Cutleriales; Desmarestiales; Ectocarpales; Laminarales; Fucales. Their occurrence as fossils is rare due to their generally soft‐bodied habit, and scientists continue to debate the identification of some finds. Only a few species of brown algae deposit significant quantities of minerals in or around their cell walls. Other algal groups, such as the red algae and green algae have a number of calcareous members, which are more likely to leave evidence in the fossil record than the soft bodies of most brown algae. Miocene fossils of a soft‐bodied brown macro algae, Julescrania, have been found well‐preserved in Monterey Formation diatomites, but few other dubiously assigned fossils, particularly of older specimens are known in the fossil record (Coyer et al., 2001). Life cycle ‐ The life cycle shows great variability from one group to another. However the life cycle of Laminaria consists of the diploid generation, that is the large kelp well known to most people. It produces sporangia from specialised microscopic structures, these divide meiotically (meiosis) before they are released. As they are haploid there are equal numbers of male and female spores (Thomas, 2002). With the exception of the Fucales all brown algae have a life cycle which consists of an alternation between haploid and diploid forms. Ecology ‐ Brown algae have adapted to a wide variety of marine ecological niches including the tidal splash zone, rock pools, the whole intertidal zone and relatively deep near shore waters. They are an important constituent of some brackish water ecosystems, and four species are restricted to life in fresh water (Lee, 2008). A large number of Phaeophyceae are intertidal or upper littoral, and they are predominantly cool and cold water organisms that benefit from nutrients in up welling cold water currents and inflows from land; Sargassum being a prominent exception to this generalisation. Brown algae growing in brackish waters are almost solely asexual (Lee, 2008). Chemistry ‐ Brown algae have a δ13C value between −‐20.8‰ – −10.5‰, in contrast with red algae and greens. This reflects their different metabolic pathways (Fletcher et al., 2004). They have cellulose walls with alginic acid; fucoidin also important in amorphous section of cell walls. A few species (of Padina) calcify with aragonite needles (Lee, 2008).
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3.4.2
Eustigmatophyceae
Eustigmatophyceae (Hibberd, 1981; Andersen et al., 1998) are a small group (7 genera; 12 species) of eukaryotic algae that includes marine, freshwater and soil‐living species (van den Hoek et al., 1995). In terms of ecology, eustigmatophyceae occur as photosynthetic autotrophs across a range of systems. Most genera live in freshwater (Fawley, 2007) or in soil, although Nannochloropsis (Guiry and Guiry, 2007) contains marine species of picophytoplankton (2 → 4 μm). Cell structure ‐ All eustigmatophyceae are unicellular, with coccoid cells and polysaccharide cell walls. They contain one or more yellow‐green chloroplasts, which contain chlorophyll a and the accessory pigments violaxanthin and β‐carotene. Eustigmatophyte zoids (gametes) possess a single or pair of flagella, originating from the apex of the cell. Unlike other heterokontophyta, eustigmatophyceae zoids do not have typical photoreceptive organelles (or eyespot); instead, an orange‐red eyespot outside of the chloroplast is located at the anterior end of the zoid. 3.4.3
Other classes
Yellow‐green algae or Xanthophyceae are an important group of heterokont algae. Most live in freshwater, but some are found in marine and soil habitats. They vary from single‐celled flagellates to simple colonial and filamentous forms. Xanthophyceae chloroplasts contain the photosynthetic pigments chlorophyll a, chlorophyll c, β‐Carotene, and the xanthophylls vaucheriaxanthin, duatoxantin and diadinoxanthin. Unlike other heterokonts, their chloroplasts do not contain fucoxanthin, which accounts for their lighter colour. Its storage polysaccharide is chrysolaminarin. Xanthophyceae cell walls are produced of cellulose and hemicellulose. They appear to be the closest relatives of the brown algae (Stace, 1991). Recent ultrastructural and molecular phylogenetic DNA (nuclear and plastid) research shows that the Mischococcales might be paraphyletic, and the Tribonematales and Botrydiales polyphyletic, and suggests two orders at most be used until the relationships within the division are sorted (Adl et al., 2005). The golden algae or Chrysophyceae are a large group of algae found mostly in freshwater. Originally they were taken to include all such forms except the diatoms and multicellular brown algae, but since then they have been divided into several different groups based on pigmentation and cell structure. They are now usually restricted to a core group of closely related forms, distinguished primarily by the structure of the flagella in motile cells, also treated as an order Chromulinales. It is possible membership will be revised further as more species are studied in detail. Most members are unicellular flagellates, with either two visible flagella, as in Ochromonas, or sometimes one, as in Chromulina. Most genera have no cell covering, some have loricae or shells. Some members are generally amoeboid, with long branching cell extensions, though they pass through flagellate stages as well. Other members are non‐motile. Cells may be naked and embedded in mucilage, such as Chrysosaccus, or coccoid and surrounded by a cell wall, as in Chrysosphaera. A few are filamentous or even parenchymatous in organization, such as Phaeoplaca.
3.5 Labyrinthulea (phylum Heterokonta) Thraustochytrids are exclusively marine heterotrophic protists that feed non‐phagotrophically as saprobes, epibionts on algae (micro and macroalgae) or more rarely as parasites of microalgae (such as Skeletonema) and animals. These unicellular eukaryotic protists are a common component of marine AquaFUELs‐ Taxonomy, Biology and Biotechnology
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microbial consortia. Whether to include or not this group among algae is still object of debate in the scientific community, but due to their phylogenetic vicinity and to the fact that products from these organism are on the market as “algal products” they are discussed in this document. Cell structure – Thraustochytriidae present an ectoplasmic net (EN) that forms a branched network of plasma membrane extensions, associated with an organelle termed the bothrosome or sagenogenetosome (sagenogen) at the periphery of the cell. The EN contributes to the increased surface area of the cell and contains hydrolytic enzymes that are surface‐bound or are secreted into the surrounding medium, helping in the digestion of organic material. The EN also attaches the cells to surfaces and, in the case of thraustochytrids, penetrates organic particles (Raghu Kumar, 2002). Their cell walls are composed of non‐cellulosic scales and contain sulphated polysaccharides, predominantly of galactose or fucose, and proteins (Leander and Porter, 2001; Raghu Kumar, 2002). Vegetative stages of thraustochytrids consist of single cells which are globose to subglobose, measuring 4 to 20 µm in diameter, mostly growing epibiontically on various substrata. Phylogeny – The Labyrinthulomycota are a group of microorganisms of uncertain taxonomic collocation. They were placed among fungi due to their functional ecology and morphology. They were then placed in the group of Oomycetes due to the presence of biflagellate zoospores. (Raghu Kumar, 1996). Cavalier Smith et al. (1994) includes the Labyrinthulomycetes in the phylum Heterokonta, based on the analysis of the 18S rRNA. The Class Labyrinthulea is then subdivided in the two sub‐classes Thraustochytridae and Labyrinthulidae. Reproduction – The Labyrinthulomycetes possess simple, asexual life cycles. Most thraustochytrids reproduce by means of zoospores which possess a long anterior, tinsel flagellum and a short, posterior, whiplash flagellum. The mode of production of zoospores varies between genera and forms the major taxonomic criterion (Raghu Kumar, 2002). The cytoplasmic contents of the mature cell, the sporangium, divide directly into zoospores in the genus Thraustochytrium. The cytoplasm escapes as one amoeboid mass, prior to zoospore division in the genus Ulkenia. The genus Schizochytrium is characerised by successive bipartition of a vegetative cell, resulting in a cluster of cells, each of which develops into a zoosporangium or zoospore. Species within the genera are primarily defined by the number of proliferating bodies and the size and shape of the sporangia.
3.6 Bacillariophyta (Diatoms) Despite the abundance and diversity of diatoms in nature, few species are cultured in aquaculture or for biotechnology relevant products (Lebeau and Robert, 2003). Further, only a handful of diatoms have been studied and often only one or a few strains without any information on intraspecific variation. There is a need to identify new diatom strains with as much positive characteristics as possible or to breed or select for improved strains. Therefore, a basic knowledge of physiology, ecology and taxonomy is important. Diatoms are the most species‐rich and productive group of eukaryotic algae. Over a comparatively short evolutionary time (50 000 and >160 000 ESTs available online. Gross composition under optimal and stressed conditions The variability of algal biochemical composition has been impressively demonstrated in C. reinhardtii during its diurnal growth cycle. During the early light period most cellular mass is protein, during the second half of the light period the alga accumulates up to 50% starch. While oil droplets in Chlamydomonas were not originally reported, recent studies revealed up to 20 % TAG under nitrogen starvation.
BIOTECHNOLOGY Clamydomonas is not being investigated for mass cultivation and biomass production, though Hu et al. (2008) have included two Chlamydomonas species in their comparative study on lipid productivity with C. applanta showing higher productivity than C. reinhardtii. However, the wide knowledge on its genetics and cell biology has allowed revealing numerous mechanisms of interest to biofuels production. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Analysis of starch deficient mutants revealed the tight link between carbon allocation to starch or lipid biosynthesis. Identification of mirna in Chlamydomonas has revealed their usefulness in upregulation of lipid biosynthesis (Maor, personal communication). Clamydomonas has the potential to produce large amounts of hydrogen because it can directly split water into hydrogen and oxygen using the enzyme hydrogenase. It was discovered that when the alga is deprived of essential sulfate salts, it no longer maintains the protein complex necessary for photosynthetic production of oxygen and instead switches to the hydrogen‐producing metabolic pathway. However, the alga can not grow in a sulfur deprived condition for long time before it needs to revert to the oxygen producing mode (Torzillo et al., 2009; Ghirardi et al., 2010). Detailed understanding of the hydrogen production process and the hydrogenase complex of Chlamydomonas have allowed creating mutants with enhanced hydrogen production capacity. Significant efforts have been undertaken to investigate expression of recombinant proteins in Chlamydomonas reinhardtii (Mayfield et al., 2003). Among the proteins expressesd successfully are antibody molecules, bovine mammary‐associated serum amyloid that may be expressed successfully both in the cytoplasm or chloroplast. (e.g. Mayfield et al., 2007). Culture media C. reinhardiii can be cultivated in a large variety of synthetic growth media such as sager‐granick, high salt (sueoka), tap, in presence or absence of acetate (Harris, 1989, p 31 on). Presence of acetate significantly enhances the growth rate under diurnal light regime. Production system Possible biohydrogen production has been investigated and modelled in a study to nrel NREL (Wade and Amos, 2004), though no actual field tests and up scaling have been performed as actual studies have been performed on the lab scale so far. For the production of recombinant proteins rather small well controlled photobioreactors are suggested rather than large outdoors cultivation systems. HIGHLIGHTS IN BIOTECHNOLOGY • • •
Early genetic work, genetic transformation and fully sequenced genome make this alga the favorite for exploring the power of genetic engineering in algal biotechnology (Harris, 1989). Major breaktroughs are algae with increased hydrogen production, reduced chlorophyll etc. Mutants with increased oil content due to impaired starch synthesis (Li et al., 2010), and application of mirna miRNA technology for enhanced oil production (Maor, personal communication). Furthermore the algae has been used to express and produce recombinant medically relevant peptides and proteins (Mayfield et al., 2003, 2007).
References Breidenbach E., Leu S., Michaels A., Boschetti A. (1990) Synthesis of EF‐Tu and distribution of tis mRNA between stroma and thylakoids during the cell cycle of Chlamydomonas reinhardii. Biochimica et Biophysica Acta 1048: 209‐216. Ghirardi M.L., Kosourov S., Maness P., Smolinski S., Seibert M. (2010) Algal Hydrogen production. In: Flickinger M.C., Anderson S. (eds.) Encyclopedia of Industrial Biotechnology: Bioprocess, Bioseparation, and Cell Technology. John Wiley & Sons, Inc., Hoboken, NJ, USA. Vol.1, pp. 184‐198. Harris E. (1989) The Chlamydomonas Sourcebook: A Comprehensive Guide to Biology and Laboratory Use.Academic Press, San Diego.
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Hu Q., Sommerfeld M., Jarvis E., Ghirardi M., Posewitz M., Seibert M., Darzins A. (2008) Microalgal triacylglycerols as feedstocks for biofuels production: perspectives and advances. The Plant Journal 54: 621‐639. Leu S., White D., Michaels A. (1990) Cell cycle‐dependent transcriptional and post‐transcriptional regulation of chloroplast gene expression inChlamydomonas reinhardtii. Biochimica et Biophysica Acta 1049: 311‐317.
Li Y., Han D., Hu G., Dauvillee D., Sommerfeld M., Ball S., Hu Q. (2010) Chlamydomonas starchless mutant defective in ADP‐glucose pyrophosphorylase hyper‐accumulates triacylglycerol. Metabolic Engineering 12: 387‐391. Mayfield S.P., Manuell A.L., Chen S., Wu J., Tran M., Siefker D., Muto M., Marin‐Navarro J. (2007) Chlamydomonas reinhardtii chloroplasts as protein factories. Current Opinion in Biotechnology 18: 126‐133. Mayfield S.P., Franklin S.E., Lerner R.A. (2003) Expression and assembly of a fully active antibody in algae. Proceedings of the National Academy of Science of the USA 100: 438‐442. Molnar A., Bassett A., Thuenemann E., Schwach F., Karkare S., Ossowski S., Weigel D., Baulcombe D. (2009) Highly specific gene silencing by artificial microRNAs in the unicellular alga Chlamydomonas reinhardtii. The Plant Journal 58: 165‐174. Torzillo G., Scoma A., Faraloni C., Ena A., Johanningmeier U. (2009) Increased hydrogen photoproduction by means of a sulfur deprived Chlamydomonas reinhardtii D1 protein mutant. International Journal of Hydrogen Energy 34: 4529‐4536.
Wade A., Amos A. (2004) Updated cost analysis of photobiological hydrogen production from Chlamydomonas reinhardtii green algae. Milestone completion report, NREL/mp‐560‐35593.
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8.1.5
Haematococcus pluvialis
Figure 20 ‐ Green and astaxanthin containing H. pluvialis cells (light microscope). D. Reinecke, BGU
SYMBOLS: B, PIV
TAXONOMY
Phylum Class Order Family Genus Species
Chlorophyta Chlorophyceae Volvocales Haematococcaceae Haematococcus Haematococcus pluvialis
Related Species There are 15 species (and infraspecific) names in the database at present, of which 7 have been flagged as currently accepted taxonomically. H. allmanii, H. buetschlii, H. capensis, H. carocellus, H. droebakensis var. fastigatus, H. droebakensis, H. grevillei, H. insignis, H. lacustris, H. murorum, H. pluvialis, H. salinus, H. sanguineus, H. thermalis, H. zimbabwiensis.
BIOLOGY Structural and morphological features Haematococcus pluvialis is a medium to large unicellular green algae (10 – 100 μm) with a large single cloroplast, two flagellae. Young cells are motile, ageing cells (palmelloids) lose their mobility. Under stress the algae accumulates the carotenoid asthaxanthin in cytoplasmic oil globules and enters a resting stage (Droop, 1954). The cell wall thickens and alters its chemical composition during maturation. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Biochemical composition of algae and aquatic biomass main constituents During steady state growth the algae is rich in protein, with less than 10% lipid content, displaying the typical green algal fatty acid composition with significant amounts of short (C16, C18) polyunsaturated fatty acids mainly in the chloroplast lipids, chlorophylls a and b and photosynthetic carotenoids such as β‐ carotene. Under stress the alga accumulates initially up to 50% starch, then initiates synthesis of triacylglycerol (TAG) accumulating to 40% of cell weight in cytoplasmic oil globules, and up to 4% of the ketocarotenoid astaxanthin (Boussiba et al., 1992, 1999). Growth kinetics and efficiencies Haematococcus is considered a slow growing alga. Productivity in outdoors systems has been reported by Huntley and Redalje (2007), implying a photosynthetic energy conversion efficiency of 3%.
BIOTECHNOLOGY Culture Media BG11 (Rippka et al., 1979), mBG11 or similar synthetic growth media. Cultivation methods Haematococcus is cultivated both in open ponds or closed bioreactors. The tubular photobioreactor facility at Qetura is the largest operating photobioreactor facility for microalgae production. Large scale outdoors cultivation in two stage mode, photobioreactor for green cells and open ponds for production of red cells was tested in Hawaii and yielded a long term growth average of 38 tons per hectare and year with 25% oil content, or 422 GJ ha‐1 year‐1 (Huntley and Redalje, 2007). Production system Production is managed as semicontinuous or batch cultivation in two stages. The first stage is a nutrient sufficient green stage that can be handled as continuous or semicontinuous culture (Boussiba et al., 1997; Huntley and Redalje, 2007), while the second stage under nutrient limitation for accumulation of astaxanthin is necessarily a batch cultivation where all the resulting biomass is harvested. One step cultivation has been proposed but not commercially deployed (Garcia‐Malea et al., 1999). Harvesting methods Haematococcus, specifically stressed resting cells settle spontaneously and can easily be harvested in large funnels or sedimentation ponds. However centrifugation yields higher recovery of biomass faster. Biomass processing The biomass is dried,dried; cells are broken by bead mills or other suitable technologies. Astaxanthin containing oil is being extracted, e. g. by supercritical CO2 extraction, and sold. Upscaling limitations Contaminations, competitors, cold or heat stress can significantly reduce productivity outdoors, and in bioreactors heating or cooling are required for maintaining satisfactory productivities. Being a freshwater microalgae, Haematococcus cultivation is frequently hampered by other, fast growing microalgae such as
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Scenedesmus or Chlorella, zooplankton or even funghi which can drastically reduce the culture performance. HIGHLIGHTS IN BIOTECHNOLOGY Astaxanthin production at the tubular bioreactor facility at Algatech Qetura (http://www.algatech.com/, Boussiba et al., 1997) may be considered the most advanced microalgal production process having reached full commercialization. This production facility has been running successfully for eight years uninterrupted under continuous incorporation of innovations for increasing productivity and product quality. References Boussiba S., Bing W., Yuan J.P., Zarka A., Chen F. (1999) Changes in pigments profile in the green alga Haematococcus pluvialis exposed to environmental stresses. Biotechnology Letters 21: 601‐604. Boussiba S., Fan L., Vonshak A. (1992) Enhancement and determination of astaxanthin accumulation in green alga Haematococcus pluvialis. Methods in Enzymology. New York: Academic Press, pp. 386‐371. Boussiba S., Vonshak A., Cohen Z., Richmond A. Ben‐Gurion University of the Negev, Israel (1997). A procedure for large‐scale production of astaxanthin from Haematococcus WO 97/28274. Droop M. (1956a) Haematococcus pluvialis and its allies. I. The Sphaerellaceae. Revue Algologique 2: 53‐70. Droop M. (1956b) Haematococcus pluvialis and its allies. II. Nomenclature in Haematococcus. Revue Algologique 3: 182‐192. Elliott A. (1934) Morphology and life history of Haematococcus pluvialis. Archiv für Protistenkunde 82L.: 250‐272. Garcia‐Malea M.C., Acien F.G., Del Rio E., Fernandez J.M., Ceron M.C., Guerrero M.G., Molina‐Grima E. (2009) Production of astaxanthin by Haematococcus pluvialis: Taking the onestep system outdoors. Biotechnology and Bioengineering 102: 651‐ 657. Hepperle D., Nozaki H., Hohenberger S., Huss V.A., Morita E., Krienitz L.. (1998) Phylogenetic position of the Phacotaceae within the Chlamydophyceaeas revealed by analysis of 18S rDNA and rbcL sequences. Journal of Molecular Evolution 47: 420‐30. Huntley M.E., Redalje D.J. (2007) CO2 mitigation and renewable oil from photsynthetic microbes: A new appraisal. Mitigation and Adaptation Strategies for Global Change 12: 573‐608. Kobayashi M., Kurimura Y., Kakizono .T, Nishio N., Tsuji Y. (1997) Morphological changes in the life cycle of the green alga Haematococcus pluvialis. Journal of Fermentation and Bioengineering 84: 94‐97. Rippka R., Deruelles J., Waterbury J.B., Herdman M., Stanier R.Y. (1979). Generic assignment, strains histories and properties of pure cultures of cyanobacteria. Journal of General Microbiology 111: 1‐61. Triki A., Maillard P., Gudin C. (1997) Gametogenesis in Haematococcus pluvialis Flotow (Volvocales, Chlorophyta). Phycologia 36: 190‐194.
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8.1.6
Dunaliella sp.
Figure 21 – Dunaliella sp. © D.J. Patterson http://starcentral.mbl.edu/msr/rawdata/viewable/dunaliel la_1097788928_esmlaw.jpg; Left:
Figure 22 ‐ Dunaliella tertiolecta Roscoff Culture Collection Strain # 6 CNRS Station Biologique de Roscoff www.sb‐ roscoff.fr/Phyto/gallery/main.php?g2_itemId=371
SYMBOLS: B, PIV
TAXONOMY
Phylum Class Order Family Genus Species
Chlorophyta Chlorophyceae Volvocales Dunaliellaceae Dunaliella
Related Species There are 28 species (and infraspecific) names in the database at present, of which 23 have been flagged as currently accepted taxonomically. D. acidophila, D. assymetica, D. baasbeckingii, D. bardawil, D. bioculata, D. carpatica, D. cordata, D. euchlora, D. gracilis, D. granulata, D. lateralis, D. maritima, D. media, D. minuta, D. parva, D. peircei, D. polymorpha, D. primolecta, D. pseudo‐salina, D. quartolecta, D. ruineniana, D. salina, D. terricola, D. tertiolecta, D. turcomanica, D. viridis var. palmelloides, D. viridis, D. viridis f. euchlora.
BIOLOGY Structural and morphological features Dunaliella is a genus of algae, specifically belonging to the Dunaliellaceae family. Dunaliella sp. are motile, unicellular, rod to ovoid shaped (9 ‐ 11 µm) green algae, Chlorophyta, which are common in marine waters. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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The genus Dunaliella has marine and halophilic representatives. Freshwater species have also been described. Dunaliella also has a very wide pH tolerance ranging from pH 1 (D. acidophila) (Gimmler et al., 1989) to pH 11 (D. salina). In fact, D. salina is one of the most environmentally tolerant eukaryotic organisms known and can cope with a salinity range from seawater (= 3% NaCl) to NaCl saturation (= 31% NaCl), and a temperature range from 38 °C (Ginzburg, 1987; Borowitzka, 1988). Dunaliella is morphologically similar to Chlamydomonas, with the main difference being the absence of a cell wall in Dunaliella. It has two flagella of equal length and a single, cup‐shaped chloroplast, which in the marine and halophilic species has a central pyrenoid (Borowitzka, 1988). D. salina is a halophile microalga and was originally identified in sea salt fields. It is different from all other green algae cells because it lacks a cell wall and is wrapped with an extremely thin elastic membrane. The lack of cell wall allows the cell to change its volume with changing osmotic pressure. In the Dead Sea it was first discovered in 1941. Another strain of interest to biotechnology is Dunaliella tertiolecta. D. tertiolecta is a marine microalga with a cell size of 10 – 12 µm. D. tertiolecta is a fast growing strain in sea water, reported to have oil yield of about 37%, though comparative growth rates and lipid productivities have not been reported. Biochemical composition In order to survive under such extreme conditions, Dunaliella synthesizes different compounds in a very high concentration. Its resistance against high salt concentrations (35%) is linked to its ability to synthesize and accumulate glycerol up to 10% of its dry weight. Due to this it can maintain shape and functions under high osmotic pressures. In order to grow in an environment of high temperature and intense sunlight, Dunaliella synthesizes beta‐carotene up to 6 of its dry weight. Studies show that the function of beta‐ carotene is to protect the algae cell from damages caused by intense radiation. Proteins accumulate up to 60% of the dry cell weight and contain most amino acids. Proteins and amino acids are important ingredients in cosmetic preparations. They are used in order to create a contaminating layer on the skin surface while nourishing the skin cells. The carbohydrates include mono‐sugars (glucose, glactose, mannose, xylose, ribose, rhamnose), di‐ sugars and 1,4 polysaccharides ‐ glucosen and starch. The sugars, especially the poly‐sugars are used as stabilizers. They thicken and give the product a smooth and gentle texture. They absorb large quantities of water and grant the product with moisturizing traits. Attached to the negative electrical charge of the poly‐ sugars are electrolytes, released in a controlled way to the skin cells. The presence of polysaccharides in the cosmetic product allows this controlled release of the active substances in the product and offers an efficient treatment of skin diseases without using substances which risk the user in side‐effects. Lipids accumulate to 6‐18% of the dry cell weight depending on growth conditions. Fatty acids include palmitic acid, 3 – trans acid hexadecanoic, linoleic acid and arachidic acid. Beta‐carotene can accumulate up to 6% of the cell's dry weight. Glycerol assembles up to 10% of the dry cell weight. Glycerol includes monogalacto glycerol, digalacto glycerol and diacyl glycerol. In addition to high level of beta‐carotene, Dunaliella contains thiamine, pyridoxine, riboflavin, nicotinic acid, biotin and tocopherol (vitamin E). β ‐carotene, produced from Dunaliella is composed of two isomers: all trans and cis‐9, in contrast to beta‐carotene produced from carrots or synthetic beta‐carotene. β‐ carotene seems to act as photo‐protective ‘sun‐screen’ to protect the chlorophyll and the cell DNA from the high irradiance which characterizes the normal habitat of D. salina. It has also been proposed that β‐ carotene also acts as a “carbon sink” to store the excess carbon produced during photosynthesis under conditions where growth is limited but photosynthetic carbon fixation must continue.
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BIOTECHNOLOGY Since establishment of a first pilot plant for Dunaliella cultivation for β‐carotene production in the USSR in 1966, the commercial cultivation of Dunaliella for the production of β‐carotene throughout the world is now one of the success stories of halophile biotechnology (Ben Amotz and Avron, 1982). Although technically the production of glycerol from Dunaliella was shown to be possible, economic feasibility is low and no biotechnological operation presently exists. β ‐carotenoid production by Dunaliella is also not competitive with other resources, so that Dunaliella production to date is limited to the nutraceuticals market. For the accumulation of high concentrations of β ‐carotene and glycerol, the algae is cultivated at extreme salinity with reported very low productivity of about 2 g m‐2 day‐1, which is not competitive for biofuels production. Production system, harvesting and processing The larger the individual ponds used to grow the algae, the smaller the productivity of the system seems to be. In D. salina the optimum salinity for growth lies between 18 and 22% NaCl whereas the optimum salinity for carotenoid production is >27% NaCl (Borowitzka et al., 1984). Thus the optimum yield of β‐ carotene per unit volume and time is achieved at about 24% NaCl. Two strategies can be applied to maximize the production of β‐carotene. One of these strategies is a two‐stage growth process in which the algae are first grown at a low salinity (= 15% NaCl) in nutrient‐rich medium to maximize biomass production, and then transferred to a high salinity, low nutrient medium in order to induce β‐carotene production (Borowitzka et al., 1984). A similar, two‐stage production process was proposed by Chen and Chi (1981) for glycerol production from Dunaliella. NBT Eilat cultivates Dunaliella in open raceway ponds in hyper saline water obtained after passages through evaporation ponds and achieves productivities of about 2 g m‐2 day‐1. A complete description of different production processes and sites, including harvesting, processing and economics of the process can be downloaded from: http://www.wind‐sea‐algae.org/wsapresentations/day1/Ami%20BenAmotz%20WSA%20April%202009.pdf Culture media The most commonly used medium for culture of Dunaliella is Modified Johnsons Medium (Borowitzka, 1988). However, these algae can also be grown in a wide range of other media including Guillard's f/2 medium (Guillard and Ryther, 1962), modified ASP medium (McLachlan and Yentsch, 1959) and enriched seawater (Rao and Chauhan, 1984). Upscaling limitations At low salinities protozoa such as the ciliate Fabrea salina and the amoeba Heteroamoeba sp. can invade the culture and very rapidly decimate the algal production. Lower salinities generally favour the growth of the non‐carotenogenic Dunaliella species (D. viridis, D. minuta and D. parva) that can overgrow D. salina and drastically reduce the β‐carotene productivity of the pond (Burford and Borowitzka, 1987). A two‐stage process requires greater capital and running costs and thus may make the process uneconomic. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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HIGHLIGHTS IN BIOTECHNOLOGY Dunaliella was the first microalga cultivated in large scale for production of a high value product (β‐ carotene) rather than bulk biomass. Its cultivation was favoured by the hypersaline medium which reduces the number of potential contaminants during long term open pond cultivation. References Ben‐Amotz A., Avron M. (1982) The potential use of Dunaliella for the production of glycerol, β‐carotene and high‐protein feed. In: San Pietro A. (ed.) Biosaline research: A look to the future. Plenum Pub. Corp., New York, pp. 207‐214. Borowitzka M.A. (1988) Algal growth media and sources of cultures. In: Borowitzka M.A,. Borowitzka L.J. (eds.) Micro‐algal Biotechnology. Cambridge University Press, Cambridge, pp. 456‐465. Borowitzka L.J., Borowitzka M.A., Moulton T.P. (1984) The mass culture of Dunaliella for fine chemicals: from laboratory to pilot plant. Hydrobiologia 116/117: 115‐121. Burford M.A., Borowitzka L.J. (1987) Competition between Dunaliella species at high salinity. Hydrobiologia 151/152: 107‐116. Chen B.J., Chi C.H. (1981) Process development and evaluation for algal glycerol production. Biotechnology and Bioengineering 23: 1267‐1287. Cifuentes A. (1992) Growth and carotenogenesis in eight strains of Dunaliella salina Teodoresco from Chile. Journal of Applied Phycology 4: 111‐118. Gimmler H, Weis U., Weiss C., Kugel H., Treffny B. (1989) Dunaliella acidophila (Kalina) Masyuk ‐ an alga with a positive membrane potential. New Phytologist 113: 175‐184. Ginzburg M.(1987) Dunaliella: a green alga adapted to salt. Advances in Botanical Research 14: 93‐183. Guillard R.R.L., Ryther J.H. (1962). Studies of marine planktonic diatoms. I. Cyclotella nana (Hustedt) and Detonula confervacea (Cleve). Canadian Journal of Microbiology 8: 229‐239. Jimenez C., Pick U. (1993) Differential reactivity of β‐carotene isomers from Dunaliella bardawil toward oxygen radicals. Plant Physiology 101: 385‐390. McLachlan J., Yentsch C.S. (1959) Observations on the growth of Dunaliella euchlora in culture. The Biological Bulletin 116: 461‐471. Rabbani S., Beyer P., Von Lintig J., Hugueney P., Kleinig H. (1998) Induced beta‐carotene synthesis driven by triacylglycerol deposition in the unicellular alga Dunaliella bardawil. Plant Physiology 116: 1239‐1248. Rao P.S.N., Chauhan V.D. (1984) On occurrence and growth of Dunaliella from India. I. Enriched seawater for mass culture of the alga. Phykos 23: 33‐37.
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8.1.7
Chlorococcum sp.
Figure 23 ‐ Chlorococcum sp. UTEX 819, D. Reinecke, BGU
SYMBOLS: H, D, E
TAXONOMY
Phylum Class Order Family Genus Species
Chlorophyta Chlorophyceae Chlorococcales Chlorococcaceae Chlorococcum
Related Species There are 76 species (and infraspecific) names in the database at present, of which 34 have been flagged as currently accepted taxonomically. C. acidum, C. aegyptiacum, C. botryoides, C. choloepodis, C. citriforme, C. costatozygotum, C. diplobionticum, C. dissectum, C. echinozygotum, C. elbense, C. elkhartiense, C. ellipsoideum, C. hypnosporum, C. infusionum, C. isabeliense, C. lobatum, C. macrostigmatum, C. minimum, C. minutum, C. novae‐angliae, C. oleofaciens, C. olivaceum, C. pamirum, C. pinguideum, C. polymorphum, C. pseudodictyosphaerium, C. pyrenoidosum, C. refringens, C. salinum, C. schizochlamys, C. schwarzii, C. submarinum, C. tatrense, C. vacuolatum.
BIOLOGY Vegetative cells solitary or in temporary groups of indefinite form, never embedded in gelatin. Cells ellipsoidal to spherical and vary in size. Cell walls smooth. The chloroplast is cup‐shaped, parietal, with or without a peripheral opening and has one or more pyrenoids. Cells uniucleate, or multinucleate just prior to zoosporogenesis. Reproduction by zoospores, aplanospores, or isogametes. Motile cells have two equal flagella and remain ellipsoidal for a time after motility ceases. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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This free‐living genus is cosmopolitan. Though primarily an edaphic alga, it has been reported from such diverse habitats as hot springs in Central Asia and soils collected in Antarctica. Aquatic, marine, and aerial isolates have been recorded.
BIOTECHNOLOGY Several strains of Chlorococcum have been tested as potential sources of astaxanthin (Zhang and Lee, 1997; Liu and Lee, 2000; Masojídek et al., 2000; Ma and Chen, 2001). Outdoor cultures to verify astaxanthin yield have been performed. An enhanced astaxanthin producing mutant was cultivated in a tubular‐loop photobioreactor consisting of two inclined panels with 34 Pyrex glass tubes each (each tube approximately 1 m long and 1.2‐cm in diameter), reaching biomass productivities up to 0.3 g L‐1 day‐1 and ketocarotenoid productivities up to 3.4 mg L‐1 day‐1 (Zhang and Lee, 1999). In a horizontal 50‐L tubular photobioreactor Masojídek et al. (2000) obatined growth rates about 0.13 h‐1, four times that of Haematococcus, but with 20 times lower secondary carotenoid content. In 10‐L bubbled tubes Tredici and co‐workers (unpublished data) obtained productivities of 0.20 g L‐1 day‐1 outdoors and 0.27 g L‐1 day‐1 indoors under contnuos illumination, with temperature reaching at mid day values up to 45 and 36 °C, respectively. In a flat‐plate photobioreactor under artificial light of 2000 μmol photons m‐2 s‐1, a ultra‐high density culture of the marine Chlorocococcum littorale reached a productivity of 380 ± 20 mg l‐1 h‐1, with 1‐cm light path length, a value 2.4 and 6.4 times higher than those obtained in the 2‐ and 4‐cm reactors. Culture denisites as high as 84 g L‐1 were reached and daily CO2 fixation rate was 16.7 g L‐1 (Hu et al., 1998). Chlorococcum has been investigated as hydrogen producer (Schnackenberg et al., 1995; Ueno et al., 1999), but hydrogen yields are much lower than those obtained with Chlamydomonas and Scenedesmus spp. (Winkler et al., 2002). Chlorococcum was also proposed for bioethanol production via dark fermentation of starch (Ueno et al., 1998; Harun and Danquah, 2011; Harun et al., 2011) and was investigated as a source of lipid for biodiesel production (Rodolfi et al., 2009; Halim et al., 2011). References Algaebase: http://www.algaebase.org/search/genus/detail/?genus_id=37477 [BIOLOGY section] Halim R., Gladman B., Danquah M.K., Webley P.A. (2011) Oil extraction from microalgae for biodiesel production. Bioresource Technology 102: 178‐185. Harun R., Danquah M.K. (2011) Influence of acid pre‐treatment on microalgal biomass for bioethanol production. Process Biochemistry 46: 304‐309. Harun R., Jason W.S.Y., Cherrington T., Danquah M.K. (2011) Exploring alkaline pre‐treatment of microalgal biomass for bioethanol production. Applied Energy: doi:10.1016/j.apenergy.2010.10.048. Hu Q., Kurano N., Kawachi M., Iwasaki I., Miyachi S. (1998) Ultrahigh‐cell‐density culture of a marine green alga Chlorococcum littorale in a flat‐plate photobioreactor. Applied Microbiology and Biotechnology 49: 655‐662. Liu B.H., Lee Y.K. (2000) Secondary carotenoids formation by the green alga Chlorococcum sp. Journal of Applied Phycology 12: 301‐ 307. Ma R.Y.N., Chen F. (2001) Enhanced production of free trans‐astaxanthin by oxidative stress in the cultures of the green microalga Chlorococcum sp. Process Biochemistry 36: 1175‐1179. Masojídek J., Torzillo G., Kopecký J., Koblížek M., Nidiaci L., Komenda J., Lukavská A., Sacchi A. (2000) Changes in chlorophyll fluorescence quenching and pigment composition in the green alga Chlorococcum sp. grown under nitrogen deficiency and salinity stress. Journal of Applied Phycology 12: 417‐426. Rodolfi L., Chini Zittelli G., Bassi N., Padovani G., Biondi N., Bonini G., Tredici M.R. (2009) Microalgae for oil: strain selection, induction of lipid synthesis, and outdoor mass cultivation in a low‐cost photobioreactor. Biotechnology and Bioengineering 102: 100‐112. Schnackenberg J., Ikemoto H., Miyachi S. (1995) Relationship between oxygen‐evolution and hydrogen‐evolution in a Chlorococcum strain with high CO2‐tolerance. Journal of Photochemistry and Photobiology B: Biology 28: 171‐174. Ueno Y., Kurano N., Miyachi S. (1998) Ethanol production by dark fermentation in the marine green alga, Chlorococcum littorale. Journal of Fermentation and Bioengineering 86: 38‐43. Ueno Y., Kurano N., Miyachi S. (1999) Purification and characterization of hydrogenase from the marine green alga, Chlorococcum littorale. FEBS Letters 443: 144‐148. Winkler M., Hemschemeier A., Gotor C., Melis A., Happe T. (2002) [Fe]‐hydrogenases in green algae: photo‐fermentation and hydrogen evolution under sulfur deprivation. International Journal of Hydrogen Energy 27: 1431‐1439.
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Zhang D.H., Lee Y.K., Ng M.L., Phang S.M. (1997) Enhanced accumulation of secondary carotenoids in a mutant of the green alga, Chlorococcum sp. Journal of Applied Phycology 9: 147‐155. Zhang D.H., Lee Y.K. (1999) Ketocarotenoid production by a mutant of Chlorococcum sp. in an outdoor tubular photobioreactor. Biotechnology Letters 21: 7‐10.
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8.1.8
Neochloris oleoabundans
Figure 24 ‐ Neochloris oleoabundans Picture from http://www.sbs.utexas.edu/utex/algaeDetail.aspx?algaeID=3623
SYMBOLS: D, PIV
TAXONOMY
Phylum Class Order Family Genus Species
Chlorophyta Chlorophyceae Chlorococcales Chlorococcaceae Neochloris Neochloris oleoabundans
Related Species There are 19 species (and infraspecific) names in the database at present, of which 11 have been flagged as currently accepted taxonomically.
BIOLOGY Structural and morphological features Neochloris oleoabundans was isolated from the top of a sand dune (2O N; 55O E) in Rub al Khali in Saudi Arabia and named by Bold and Chantanachat (Chantanachat and Bold, 1962). Vegetative cells are described as being between 6 and 25 μm, with a cup shaped parietal chloroplast. Vegetative cells with smaller diameter (3.5 μm) however are most common with the strain UTEX 1185, which originates from the original AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Bold and Chantnachat isolate. After cell division, one pyrenoid is present in the chloroplast and later in the cell cycle it divides into two. The cells are uninucleate. Sexual reproduction has not been observed (Chantanachat and Bold, 1962); N.oleoabundans reproduces asexually through formation of zoospores or aplanospores. Zoospores are biflagellate (identical length of flagella) and between 2 and 3.5 μm in width and 3.6 and 4.5 μm in length. Zoospores form vegetative sphaerical cells after a short period of time and are not often observed in laboratory cultures while they appear with varying frequency in outdoor mass cultures. Vegetative cells may be up to 22 μm in diameter and may contain visible oil droplets in the cytoplasm. Aplanospores are formed by incomplete separation of daughter cells: the wall protoplasts (wall less and non‐motile cells) are retained within the cell wall of the original mother cell (termed sporangium). Aplanospores are about the size of zoospores and may accumulate to considerable numbers before they are liberated by rupture of the cell wall of the sporangium. Biochemical composition of algae and aquatic biomass main constituents Biochemical composition in Neochloris varies with growth stage which, in a more “modern”concept may be considered an effect of growth limitation by either light or nutrient factors. In Table 6 it may be noted that protein content significantly decreases in the stationary phase while lipid and carbohydrate content increase from exponential to stationary phase. InTable 7, it may be noted that neither total fatty acid‐, unsaturated fatty acid‐ or sterol percentage of lipids are affected by growth stage to any appreciable extent. Table 6 ‐ Example of biochemical composition through growth stages given in percent dry weight. Source of growth limitation ‐2 ‐1 not indicated. Algae were cultivated in 100 ml test tubes at low irradiation (60 μm m sec ). After Gatenby et al. (2003).
Constituent ( % DW) Exponential Protein 54 Carbohydrate 8 Lipid
Late exponential 63 10
Stationary 44 18
Late stationary 18 40
22
35
36
19
Table 7 ‐ Example of lipid composition through growth stages given in percent lipid (or, for sterols, in ‰ lipid). Conditions as in table 6. After Gatenby et al. (2003).
Constituent ( % lipid) Exponential Fatty acids 32 Unsaturated FA 89 Sterols (‰ lipid)
Late exponential 45 85
Stationary 31 88
Late stationary 54 80
9.6
5.1
4.5
5.2
Gross composition under optimal and stressed conditions Published studies of the effect of stress conditions are all applying nitrogen limitation as stress factor. A single study (Pruvost et al., 2011) investigates the effect of stress on gross composition of Neochloris, other studies published so far, focus only on lipid proportions. The findings are summarized in Table 8.
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Table 8 ‐ Changes in gross composition of Neochloris through nutrient starvation. Data from artificially illuminated PBR’s.
System Unit/fraction
Flat plate PBR, 1 and 130 L Not indic.
Light μmol ‐2 photons m s‐1 or as otherwise indicated 300 ‐ 500 3 klux
1 L bubble columns
Not indic.
90 ‐ 260 mm bubble columns
160 ‐ 200
Temp o
Change in composition from nutrient replete to deplete stage. Lipid Protein Carbo‐ hydrate
C
30
20% → 30‐35 % 28 ‐31 → 35 – 55 % 34 34 % 30
12.5 % → 24.9 %
70 % → 20 %
20 % → 40 %
Starvation method
Reference
4 days nitrate starvation To visible lipid droplet accumulation 6‐7 days nitrate starvation 7 days on nitrogen free medium
(Pruvost et al., 2011)
(Tornabene et al., 1983) (Li et al., 2008) (Kawata et al. ,1998) (Neochloris sp.)
Photosynthetic efficiency and productivity From data in artificially illuminated (continuous) photobioreactors, Neochloris appears to exhibit slow to moderate growth rates (Table 9). However, much higher transient growth rates have been experienced in turbidostats at 30 °C (unpublished data, N.H. Norsker) and may be due to mixotrophic growth on internal lipid pools. No studies have yet reported figures on photosynthetic efficiency in outdoor cultivation. It has only been possible to identify one study of cultivation of Neochloris in outdoor systems. The growth rate and lipid productivity are reported in Table 10. Table 9 ‐ Growth rate and lipid productivity obtained with Neochloris under different conditions. All refer to artificially illuminated systems.
150
30
replete
1.0
0.495
37.66
1 L flat plate reactor
270
25
deplete replete
0.2
0.41
14.42 126
deplete
0.5
65
50 L plastic sleeves 1 L bubble columns 10 cm tubular horizontal PBR 90‐260 mm bubble columns
150
25
?
0.92
0.07
4
not indicated 200
30
deplete
2.4
133
23 ‐ 25
deplete
0.3
16.5
replete
0.9
8.9
replete
0.72
deplete
0 – 0.43
160 ‐ 200
30
Reference
Lipid Product. (mg L‐1 day‐1)
Growth rate (day‐1)
Biomass range (g DW L‐1)
N status (repl/depl)
Temp (°C)
Irradiation (μmol photons m‐2 s‐1)
System description 1 L bubble column.
(Gouveia et al., 2009) (Pruvost et al., 2009) (da Silva et al., 2009) (Li et al., 2008) (Levine et al., 2011)
(Kawata et al., 1998) (Neochloris sp.)
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Table 10 ‐ Growth rate and lipid productivity of Neochloris in outdoor cultivation.
System description
Irradiation Biomass Growth N status (μmol range ‐1 rate ‐1 (repl/depl) photons m‐2 (g DW L ) (day ) ‐1 s ) Outdoor natural 0.02 – replete 0.18 raceway pond daylight 2.78 (batch)
Lipid productivity (mg L‐1 day‐1) 1.6 – 4.8
Reference
(da Silva et al., 2009)
In all the studies published so‐far on Neochloris oleoabundans, the strain UTEX 1185 (the original isolate) has been applied. In only one study included in the present material (Kawata et al. 1998), the strain is not UTEX 1185 but a local strain (Neochloris sp.). There is thus insufficient material to discriminate between different laboratory strains in terms of productivity and growth rate.
BIOTECHNOLOGY Culture Media Only photoautotrophic growth of Neochloris oleoabundans has so‐far been demonstrated. The culture media used are repoted in Table 11. Table 11 ‐ Media used for N. oleoabundans by the cited references.
Reference media
Other media
BBM
3NBBM
3N3SBBM
Soil extract
B12
Other
Reference
x
x
(MD) inorganic basal medium
x x
x
Inorganic basal medium
?
x x
x
x
(modified for nutrient starvation)
(Gatenby et al., 2003) (Band et al. 1992) (Tornabene et al., 1983) (Li et al., 2008) (Archibald and Smith, 1987) (Pruvost et al., 2009) (Gouveia et al., 2009) (da Silva et al., 2009) (Pruvost et al., 2011)
x
x
x
(Levine et al., 2011)
x
x
Modified: +boron + vitamins, N‐ source) Modified: no nitrogen, + B12 Modified
x
Modified Fitzgerald
(Beal et al., 2010) (Wahal and Viamajala, 2010) (Kawata et al., 1998)
Bristol medium
Other medium
Special micro nutrient additives
A single experiment with cultivation of Neochloris in wastewater from anaerobically digested manure is reported (Levine et al., 2011): while Neochloris in a horizontal chamber, irradiated at 200 μmol photons m‐2 s‐1 on nitrate exhibited a specific growth rate of 0.75 day‐1, a similar setup resulted in a growth rate of about 0.24 day‐1 on diluted wastewater (growth rates estimated from published graphs). AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Cultivation methods Only one experiment cultivating Neochloris in open systems has been reported. The raceway area was 2.5 m2, the depth of 15 cm and it was agitated with a paddlewheel to create a flow velocity of 25 cm s‐1. The culture was diluted after 20 and 36 days of cultivation (da Silva et al., 2009). Specific growth rate during the first 20 days was 0.18 day‐1. Other systems have been used for indoor use and among these: • bubble columns of various diameter up to 26 cm (Kawata et al. 1998; Li et al. 2008; Gouveia et al. 2009) • flat panel reactors, 1 ‐130 L (Pruvost et al., 2011) • shaken flasks (Wahal and Viamajala, 2010) • plastic bags (da Silva et al., 2009; Levine et al., 2011) Production systems At present time (2010), Neochloris has not reached a state of commercial production. Neither has any information about testing of production systems in pilot scale been published. Harvesting methods No studies of harvesting methods for Neochloris have yet been published. Speculating about harvest methods, it is worth noting that the cell size of Neochloris oleoabundans is rather small (average diameter assuming sphaerical cells 3 µm for vegetatively growing cultures) and high energy input for separation by, for instance, centrifugation could be assumed. However, it is generally observed that Neochloris sediments rather readily in culture flasks and may therefore be suitable for flocculation. Flotation or foam separation are other potentially attractive harvesting technologies as Neochloris cultures are prone to foaming. Biomass processing There are no published studies on processing Neochloris biomass for any purpose. For potential use for biodiesel purposes, it is worth noting that a substantial part of the lipid fraction may be constituted by triacylglycerols: 80 % with lipids constituting 35 – 54 % of the biomass (Tornabene et al., 1983). A recent study of oil formation optimization demonstrated a TAG yield of 50 % of total lipids or 18% of DW (Pruvost et al., 2009). Relatively high specific TAG productivities by Neochloris is one of the main motives for focussing on Neochloris for biodiesel production purposes. Scaling up limitation Production of Neochloris for biodiesel is here considered only for photoautotrophic production methods. Conversion of carbohydrate sources to biodiesel by heterotrophic production is being considered with other algae species, but a heterotrophic growth potential of Neochloris has not been established. Due to the lack of experience with outdoor cultivation of Neochloris, it is very difficult to pin point limitations to scale up. Two studies have dealt with outdoor production. In da Silva et al. (2009) production of Neochloris in a 2.5 m 2 raceway pond resulted in high biomass densities (2.8 g DW L‐1) at the end of a 20 days batch growth run with specific growth rate of 0.18 day‐1 . Lipid productivity at the peak was 4.8 g m‐2 day‐1 (July, Portugal) which must be considered promising if such rates can be sustained over longer periods. It is likely that the lipid productivity can be enhanced by reducing the biomass density in the pond as high average light intensities are required to obtain high lipid productivity and the other design parameter, culture depth which determines the average light intensity in the culture, cannot be reduced in raceway ponds for AquaFUELs‐ Taxonomy, Biology and Biotechnology
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practical reasons. This will, on the other hand, add to the harvesting costs and may add to the vulnerability of the culture to contamination by other algae or protozoans and zooplankton. In Levine et al. (2011) the use of wastewater from biogas production as fertilizer was studued, but overgrowth by other algae was encountered and closed reactors were suggested for the production. For these reasons, the development of photobioreactors with low installation and operating costs is believed to be the most promising technologies for a scale up strategy.
References Arce G., Bold H.C. (1958) Some Chlorophyceae from Cuban soils. American Journal of Botany 45: 492‐503. Archibald P.A., Smith V.J. (1987) Notes on variation on physiological attributes between aquatic and edaphic species of the chlorophycean algal genus Neochloris. Transactions of the American Microscopical Society 106: 179‐182. Band C.J., Arredondovega B.O., Vazquezduhalt R., Greppin H. (1992) Effect of a salt‐osmotic upshock on the edaphci microalga Neochloris oleoabundans. Plant Cell and Environment 15: 129‐133. Beal C.M., Webber M.E., Ruoff R.S., Hebner R.E. (2010) Lipid analysis of Neochloris oleaoabundans by liquid state NMR. Biotechnology and Bioengineering 106: 573‐583. Chantanachat S., Bold H.C. (1962) Phycological Studies. II. Some algae from arid soils. University of Texas Publication (6218). da Silva T., Reis A., Medeiros R., Oliveira A., Gouveia L. (2009) Oil production towards biofuel from autotrophic microalgae semicontinuous cultivations monitorized by flow cytometry. Applied Biochemistry and Biotechnology 159: 568‐578. Gatenby C.M., Orcutt D.M., Kreeger D.A., Parker B.C., Jones V.A., Neves R.J. (2003) Biochemical composition of three algal species proposed as food for captive freshwater mussels. Journal of Applied Phycology 15: 1‐11. Gouveia L., Marques A., da Silva T., Reis A. (2009) Neochloris oleoabundans UTEX #1185: a suitable renewable lipid source for biofuel production. Journal of Industrial Microbiology and Biotechnology 36: 821‐826. Kawata M., Nanba M., Matsukawa R., Chihara M., Karube I. (1998) Isolation and characterization of a green alga Neochloris sp. for CO2 fixation. Studies in Surface Science and Catalysis 114: 637‐640. Levine R.B., Costanza‐Robinson M.S., Spatafora G.A. (2011) Neochloris oleoabundans grown on anaerobically digested dairy manure for concomitant nutrient removal and biodiesel feedstock production. Biomass and Bioenergy 35: 40‐49. Li Y., Horsman M., Wang B., Wu N., Lan C. (2008) Effects of nitrogen sources on cell growth and lipid accumulation of green alga Neochloris oleoabundans. Applied Microbiology and Biotechnology 81: 629‐636. Pruvost J., Van Vooren G., Cogne G., Legrand J. (2009) Investigation of biomass and lipids production with Neochloris oleoabundans in photobioreactor. Bioresource Technology 100: 5988‐5995. Pruvost J., Van Vooren G., Le Gouic B., Couzinet‐Mossion A., Legrand J. (2011) Systematic investigation of biomass and lipid productivity by microalgae in photobioreactors for biodiesel application. Bioresource Technology 102: 150‐158. Starr R.C. (1955) A comparative study of Chlorococcum meneghini and other spherical, zoospore‐producing genera of the Chlorococcales. Indiana University Publications Science 20: 1‐111. Starr R.C. (1978) Culture collection of algae at the University‐of‐Texas at Austin. Journal of Phycology 14: 47‐100. Tornabene T.G., Holzer G., Lien S., Burris N. (1983) Lipid composition of the nitrogen starved green alga Neochloris oleoabundans. Enzyme and Microbial Technology 5: 435‐440. Wahal S., Viamajala S. (2010) Maximizing algal growth in batch reactors using sequential change in light intensity. Applied Biochemistry and Biotechnology 161: 511‐522.
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8.1.9
Scenedesmus sp.
Figure 25 ‐ Scenedesmus sp. Local isolate Sde Boker, Israel D. Reinecke, BGU
SYMBOLS: B, D, E, PIV
TAXONOMY
Phylum Class Order Family Genus Species
Chlorophyta Chlorophyceae Sphaeropleales Scenedesmaceae Scenedesmus
Related Species There are 418 species (and infraspecific) names in the database at present, of which 87 have been flagged as currently accepted taxonomically.
BIOLOGY Scenedesmus is a freshwater medium to large size unicellular green alga often appearing in tetrads with four to 16 elongated cells connected (cenobia), although cells can also appear as individual in oval form. Scenedesmus is versatile in production of oil and secondary metabolites such as carotenoids. Multiple species produce different carotenoids during logarithmic growth or stress, though cellular levels never exceed 1% of dry weight. It is relevant that major carotenoid is usually lutein. Exponentially growing cells have high protein content (up to 50%), less than 10% lipids, the rest variable amounts of starch and cell wall components. Under nutrient stress Scenedesmus species can accumulate high amounts of storage lipids (TAG) in cytoplasmic oil globules. Fatty acid composition under logarithmic growth includes significant amounts of short polyunsaturated fatty acids such as gamma linolenic and stearidonic acid. Under stress, TAG accumulates mostly C16 and C18 unsaturated and some monounsaturated fatty acids.
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Growth kinetics and efficiencies Scenedesmus is among the most vigorously growing green algae and outcompetes most other algal species under high nutrient conditions, e. g. in wastewater. It can cause serious contamination problems when cultivating slower growing algae such as Haematococcus. It is among the faster growing and highest oil producing strains tested by Rodolfi et al. (2009) or Hu et al. (2008). Growth rates and productivity in laboratory main groups Maximum specific growth rates as higher than 0.12 h‐1 has been reported for Scenedesmus sp. Lower specific growth rates, of 0.04 h‐1, has been reported for Scenedesmus obliquus. The most relevant aspect of this strain is its tolerance to high temperature, including 40 °C, its maximum growth rate being obtained at temperatures of 30‐35 °C. At laboratory scale biomass productivities of 0.9 g∙L‐1∙day‐1 has been reported for Scenedesmus almeriensis. High productivity requires the use of high irradiance, but Scenedesmus has demonstrate to be resistant to irradiances higher than 1700 μmol photons∙m‐2∙s‐1 without photoinhibition. Scenedesmus is also tolerant to impulsion using centrifugal pumps, in addition to aeration, no mechanical damage being reported by this phenomenon. Concerning pH, Scenedesmus tolerates wide ranges of pH, from 5 to 10, although optimal pH is in the range of 7.5‐8.0. It is particularly relevant the tolerance of Scenedesmus to alkaline pH for wastewater and flue flue gas depuration. Photosynthetic efficiency and productivity outdoors Due to the fact that Scenedesmus is highly robust and fast growing, it has been produced outdoors using both open systems and closed photobioreactors. In open raceways, biomass productivities higher than 0.5 g∙L‐1∙day‐1 has been reported, while biomass productivities up to 1.2 g∙L‐1∙day‐1 have been obtained using closed photobioreactors. Overall mean annual productivity of 0.6 g∙L‐1∙day‐1 has been obtained in pilot scale tubular photobioreactors (30 m3) with Scenedesmus almeriensis. This value is about 3% solar efficiency. Solar efficiencies from 1‐3% has have been reported for Scenedesmus. Gross composition under optimal and stressed conditions Under optimal growth conditions the major component of Scenedesmus biomass are proteins, 40‐50% d.wt., next being carbohydrates. The lipid content under adequate growth conditions is lower than 15%, woith maximum values of 10% corresponding to fatty acids. It is relevant that under optimal conditions the carotenoid content, especially lutein, increases up to 1% in some species. Scenedesmus cells can be stressed by nutrient depletion, nitrogen or phosphorous, which triggers accumulation of lipids. However, no lipid content higher than 30% are obtained in these conditions.
BIOTECHNOLOGY Culture Media Scenedesmus grows well in modified BG11 medium, but also in other growth media, in nutrient rich wastewater etc, and apparently may also exploit organic molecules for photoheterotrophic growth. It has been cultivated using commercial fertilizers outdoors, and in brackish water. It is tolerant to the use of nitrate, ammonia or urea as nitrogen source, thus Scenedesmus is adequate for removal of inorganic nitrogen from effluents. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Cultivation methods Scenedesmus can be cultivated in either discontinuous (batch) or continuous mode. In discontinuous mode the productivity is lower, thus usually semi‐continuous operation being utilized. No large variation in the composition of the biomass is observed when cultivated in different modes. In continuous mode the optimal dilution rate is in the range from 0.3‐0.4 day‐1. Production system Scenedesmus are not damaged by aeration or impulsion centrifugal pumps, thus it has been cultivated in most of culture systems developed, including open and closed photobioreactors. Fouling is an important issue when cultivating this strain. Under adverse conditions, but also under optimal conditions, cells of Scenedesmus fix on the reactor’s walls, reducing light availability inside the culture. When closed reactors are used it is obligatory to implement self‐cleaning systems. This is not necessary when using open raceways. Harvesting methods Although Scenedesmus cells are larger than other microalgae cells, their settling velocity by gravity is low, in the range of 10‐6 m∙s‐1. Thus, Scenedesmus cells cannot be harvested by natural sedimentation. It is necessary the use of centrifugation or filtration operations. Cells of Scenedesmus are easily harvested by centrifugation: a slurry containing 15% d.wt is obtained under continuous operation, and a paste containing 30% d.wt. is obtained under discontinuous operation. Although filtration can be performed, it is not usually carried out. Natural sedimentation can be performed by previous flocculation. The use of anionic polyelectrolyte at doses below 0.1 mg/L has been demonstrated to be useful to pre‐concentrate the culture by twenty times. Biomass processing Biomass of Scenedesmus can be used as biomass for feeding animals or fishes, mainly for its high protein content. The fatty acid profile is not highly valuable because no PUFAs are present. The only especially valuable components present in the biomass of Scenedesmus are carotenoids. Lutein contents up to 1% d.wt. has have been reported, it being useful as nutraceutical for human and animals. Lutein can be extracted using organic solvents, although new processes using supercritical fluids or ethanol‐water mixtures have been proposed. Scaling up limitation No limitations are directly related with the culture of Scenedesmus, different than the production of other strains. For scale‐up purposes the tolerance of this strain to high temperatures, up to 45 °C, is highly relevant. HIGHLIGHTS IN BIOTECHNOLOGY Scenedesmus is highly robust and has been used successfully used to treat such problematic waste effluents as olive mill wastewater, biogas or municipal land fill effluents which are toxic to most microorganisms including bacteria. A biotechnology for production and purification of lutein from Scenedesmus has been developed based on cultivation in tubular photobioreactors and extraction and purification of the carotenoid from cellular lysate, though the technology is not currently commercially applied. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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References Abeliovich A., Dikbuck S. (1977) Factors affecting infection of Scenedesmus obliquus by a Chytridium sp. in sewage oxidation ponds. Applied Environmental Microbiology 34: 832‐836. Allard B., Templier J. (2000) Comparison of neutral lipid profile of various trilaminar outer cell wall (TLS)‐containing microalgae with emphasis on algaenan occurrence. Phytochemistry 54: 369‐380. Benemann J.R., Koopman B.L., Murry M., Weissman J.C., Eisenberg D.M., Oswald W.J. (1977) Species control in large‐scale algal biomass production. Final report. United States. Blokker P., van den Ende H., de Leeuw J.W., Versteegh G.J.M., Damste J.S.S. (2006) Chemical fingerprinting of algaenans using RuO4 degradation. Organic Geochemistry 37: 871‐881. Buchheim M.A., Michalopulos E.A., Buchheim J.A. (2001) Phylogeny of the Chlorophyceae with special reference to the Sphaeropleales: a study of 18S and 26S rDNA data. Journal of Phycology 37: 819‐835. Cepak V., Pribyl P. (2006) The effect of colour light on production of zooids in 10 strains of the green chlorococcal alga Scenedesmus obliquus. Czech Phycology 6: 127‐133. Cepak V., Pribyl P., Vitova M., Zachleder V. (2007) The nucleocytosolic and chloroplast cycle in the green chlorococcal alga Scenedesmus obliquus (Chlorophyceae, Chlorococcales) grown under various temperatures. Phycologia 46: 263‐269. Czygan F. (1968) Secondary carotenoids in green algae II. Studies on biogenesis. Archiv für Mikrobiologie 62: 209‐236. de Morais M.G., Costa J.A.V. (2007) Biofixation of carbon dioxide by Spirulina sp. and Scenedesmus obliquus cultivated in a three‐ stage serial tubular photobioreactor. Journal of Biotechnology 129: 439‐445. Dean A.P., Sigee D.C., Estrada B., Pittman J.K. (2010) Using FTIR spectroscopy for rapid determination of lipid accumulation in response to nitrogen limitation in freshwater microalgae. Bioresource Technology 101: 4499‐4507. Disch A., Schwender J., Muller C., Lichtenthaler H.K., Rohmer M. (1998) Distribution of the mevalonate and glyceraldehyde phosphate/pyruvate pathways for isoprenoid biosynthesis in unicellular algae and the cyanobacterium Synechocystis PCC 6714. Biochemical Journal 333: 381‐388. Gonzalez Lopez CV, Ceron Garcia MdC, Fernandez FG, Segovia Bustos C, Chisti Y, Fernandez Sevilla JM. 2010. Protein measurements of microalgal and cyanobacterial biomass. Bioresource Technology 101(19):7587‐7591. Greger M., Johansson M. (2004) Aggregation effects due to aluminum adsorption to cell walls of the unicellular green alga Scenedesmus obtusiusculus. Phycological Research 52: 53‐58. Grewe C., Menge .S, Griehl C. (2007) Enantioselective separation of all‐E‐astaxanthin and its determination in microbial sources. Journal of Chromatography A 1166: 97‐100. Gurbuz F., Ciftci H., Akcil A. (2008) Biodegradation of cyanide containing effluents by Scenedesmus obliquus. Journal of Hazardous Materials 162: 74‐79. Gutman J., Zarka A., Boussiba S. (2009) The host‐range of Paraphysoderma sedebokerensis, a chytrid that infects Haematococcus pluvialis. European Journal of Phycology 44: 509 ‐ 514. Hanagata N., Dubinsky Z. (1999) Secondary carotenoid accumulation in Scenedesmus komarekii (Chlorophycea, Chlorophyta). Journal of Phycology 35: 960‐966. Heide H., Kalisz H.M., Follmann H. (2004) The oxygen evolving enhancer protein 1 (OEE) of photosystem II in green algae exhibits thioredoxin activity. Journal of Plant Physiology 161: 139‐149. Ho S.H., Chen W.M., Chang J.S. (2010) Scenedesmus obliquus CNW‐N as a potential candidate for CO2 mitigation and biodiesel production. Bioresource Technology 101: 8725‐8730. Hu Q. ,Sommerfeld M. (2004). Selection of high performance microalgae for bioremediation of nitrate‐contaminated roundwater. Technical Report for Grant Number. School of Life Sciences. Arizona State University. Humbeck K. (1990) Light‐dependent carotenoid biosynthesis in mutant C‐6D of Scenedesmus obliquus. Photochemistry and Photobiology 51: 113‐118. Kim M.K., Park J.W., Park C.S., Kim S.J., Jeune K.H., Chang M.U., Acreman J. (2006) Enhanced production of Scenedesmus spp. (green microalgae) using a new medium containing fermented swine wastewater. Bioresource Technology 98: 2220‐2228. Krienitz L., Wirth M. (2006) The high content of polyunsaturated fatty acids in Nannochloropsis limnetica (Eustigmatophyceae) and its implication for food web interactions, freshwater aquaculture and biotechnology. Limnologica ‐ Ecology and Management of Inland Waters 36: 204‐210. Kuck U., Jekosch K., Holzamer P. (2000) DNA sequence analysis of the complete mitochondrial genome of the green alga Scenedesmus obliquus: evidence for UAG being a leucine and UCA being a non‐sense codon. Gene 253: 13‐18. Lee J.Y., Yoo C., Jun S.Y., Ahn C.Y., Oh H.M. (2010) Comparison of several methods for effective lipid extraction from microalgae. Bioresource Technology 101 Suppl S1: S75‐S77. Lopez‐Rodas V., Agrelo M., Carrillo E., Ferrero L.M., Larrauri A., Martin‐Otero L., Costas E. (2001) Resistance of microalgae to modern water contaminants as the result of rare spontaneous mutations. European Journal of Phycology 36: 179‐190. Lurling M. (2006) Effects of a surfactant (FFD‐6) on Scenedesmus morphology and growth under different nutrient conditions. Chemosphere 62: 1351‐1358.
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Ma J.Y., Xu L.G., Wang S.F., Zheng R.Q., Jin S.H., Huang S.Q., Huang Y.J. (2002) Toxicity of 40 herbicides to the green alga Chlorella vulgaris. Ecotoxicology and Environmental Safety 51: 128‐132. Mallick N., Mohn F.H., Soeder C.J., Grobbelaar J.U. (2002) Ameliorative role of nitric oxide on H2O2 toxicity to a chlorophycean alga Scenedesmus obliquus. Journal of General and Applied Microbiology 48: 1‐7. Masojidek J., Torzillo G., Koblizek M., Kopecky J., Bernardini P., Sacchi A., Komenda J. (1999) Photoadaptation of two members of the Chlorophyta Scenedesmus and Chlorella in laboratory and outdoor cultures: changes in chlorophyll fluorescence quenching and the xanthophylls cycle. Planta 209: 126‐135. Matsunaga T., Matsumoto M., Maeda Y., Sugiyama H,. Sato R., Tanaka T. (2009) Characterization of marine microalga, Scenedesmus sp. strain JPCC GA0024 toward biofuel production. Biotechnology Letters 31: 1367‐1372. Ordog V., Stirk W., Lenobel R., Bancirova M., Strnad M., van Staden J., Szigeti J., Nemeth L. (2004) Screening microalgae for some potentially useful agricultural and pharmaceutical secondary metabolites. Journal of Applied Phycology 16: 309‐314. Orosa M., Torres E., Fidalgo P., Abalde J. (2000) Production and analysis of secondary carotenoids in green algae. Journal of Applied Phycology 12: 553‐556. Proctor V. (1957) Studies of algal antibiosis using Haematococcus and Chlamydomonas. Limnology and Oceanography 2: 125‐139. Proctor V. (1957) Some controling factors in the distribution of Haematococcus pluvialis. Ecology 38: 457‐462. Qiang L., Junda L. (2010) Effects of nitrogen source and concentration on biomass and oil production of a Scenedesmus rubescens like microalga. Bioresource Technology 102: 1615‐1622. Qin S., Liu G.X., Hu Z.Y. (2008) The accumulation and metabolism of astaxanthin in Scenedesmus obliquus (Chlorophyceae). Process Biochemistry 43: 795‐802. Radmer R., Behrens P., Arnett K. (1987) Analysis of the productivity of a continuous algal culture system. Biotechnology and Bioenginering 29: 488‐492. Rausch T. (1981) The estimation of micro‐algal protein content and its meaning to the evaluation of algal biomass I. Comparison of methods for extracting protein. Hydrobiologia 78: 237‐251. Sánchez J.F., Fernández‐Sevilla J.M., Acién F. G., Cerón M.C., Pérez‐Parra J., Molina‐Grima E. (2008) Biomass and lutein productivity of Scenedesmus almeriensis: influence of irradiance, dilution rate and temperature. Applied Microbiology and Biotechnology 79: 719–729. Tripathi U., Sarada R., Ravishankar G.A. (2001) A culture method for microalgal forms using two‐tier vessel providing carbon dioxide environment: Studies on growth and carotenoid production. World Journal of Microbiology & Biotechnology 17: 325‐329. Vandamme D., Foubert I., Meesschaert B., Muylaert K. (2010) Flocculation of microalgae using cationic starch. Journal of Applied Phycology 22: 525‐530. Voltolina D., Cordero B, Nievesc M., Soto L.P. (1998) Growth of Scenedesmus sp. in artificial wastewater. Bioresource Technology. 68: 265‐268. Xia J.R., Gao K.S. (2002) Effects of CO2 enrichment on microstructure and ultrastructure of two species of freshwater green algae. Acta Botanica Sinica 44: 527‐531. Xin L., Hong‐ying H., Jia Y. (2010) Lipid accumulation and nutrient removal properties of a newly‐isolated freshwater microalga, Scenedesmus sp. LX1, growing in secondary effluent. New Biotechnology 27: 59‐63. Xin L., Hong‐ying H., Jia Y., Yin‐hu W. (2010) Enhancement effect of ethyl‐2‐methyl acetoacetate on triacylglycerols production by a freshwater microalga, Scenedesmus sp. LX1. Bioresource Technology 101: 9819‐9821. Yang Y., Gao K.S. (2003) Effects of CO2 concentrations on the freshwater microalgae, Chlamydomonas reinhardtii, Chlorella pyrenoidosa and Scenedesmus obliquus (Chlorophyta). Journal of Applied Phycology 15: 379‐389. Yoo C., Jun S. Y., Lee J.Y., Ahn C.Y., Oh H.M. (2010) Selection of microalgae for lipid production under high levels carbon dioxide. Bioresource Technology 101 Suppl. 1: S71‐S74.
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8.1.10 Desmodesmus sp.
Figure 26 ‐ Desmodemus sp. Picture from UNIFI
SYMBOLS: H, D, E
TAXONOMY
Phylum Class Order Family Genus Species
Chlorophyta Chlorophyceae Sphaeropleales Scenedesmaceae Desmodesmus
Related Species There are 72 species (and infraspecific) names in the database at present, of which 16 have been flagged as currently accepted taxonomically. D. communis, D. costato‐granulatus, D. bicellularis, D. serratus, D. denticulatus, D. lefevrei, D. arthrodesmiformis, Desmodesmus sp. Hegewald 1987‐51, D. subspicatus, D. hystrix, D. opoliensis, D. pannonicus, D. perforatus, D. pirkollei, Desmodesmus sp. CL1, D. maximus, D. tropicus, D. komarekii, D. multivariabilis, D. pleiomorphus, D. fennicus, D. armatus.
BIOLOGY Desmodesmus used to be the most species‐rich subgenus of Scenedesmus, but it was given genus status based on 18S and ITS2 rDNA phylogenies. The large genetic distance between the two subgenera and their clear distinct cell wall ultrastructure supported retention of the Scenedesmus Meyen for non‐spiny organisms and formation of a genus Desmodesmus (Chodat) An, Friedl et Hegewald for those which could AquaFUELs‐ Taxonomy, Biology and Biotechnology
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bear spines. Desmodesmus appears as unicells or coenobia 2–4–8–16‐celled, with long axes of cells parallel, laterally adjoined, and arranged in a single linear or alternating series. Cells are ellipsoidal to ovoid, and spines usually are present on the terminal cells and/or medial cells, but may be entirely absent. The cell wall may have ridges, warts, or nets. The chloroplast is parietal, usually with one pyrenoid. Desmodesmus is an extremely common genus, occasionally abundant, found in the phytoplankton of ponds and lakes. They are cosmopolitans and able to withstand harsh conditions, such as periods with strong grazing pressure. The relatively low number of studies with Desmodesmus can be explained from investigators still being unaware of the division of the old genus Scenedesmus into the new genera Scenedesmus and Desmodesmus.
BIOTECHNOLOGY Little scientific literature is available on Desmodesmus, although its application are similar to that reported for Scenedesmus. Part of the literature available about Scenedesmus actually refers to Desmodesmus. Virginia Coastal Energy Research Consortium cultivated a Desmodemus strain for biodiesel production studies in a 0.4 ha open pond during about a year with low and variable biomass concentrations and quite low lipid yields (Stubbins, 2009). References Algaebase: http://www.algaebase.org/search/genus/detail/?genus_id=45397 [BIOLOGY section] Lürling M. (2003) Phenotypic plasticity in the green algae Desmodesmus and Scenedesmus with special reference to the induction of defensive morphology. Annales de Limnologie ‐ International Journal of Limnology 39: 85‐101. [BIOLOGY section] Shubert L.E. (2003) Nonmotile coccoid and colonial green algae. In: Wehr J.D., Sheath R.G. (eds.) Freshwater Algae of North America: Ecology and Classification. Elsevier Science, pp. 253‐309. [BIOLOGY section] Stubbins A. (2009) Virginia Coastal Energy Research Consortium Final Report: Algal Biodiesel Studies, July 2007 to September 2009. Vanormelingen P., Hegewald E., Braband A., Kitschke M., Friedl T., Sabbe K., Vyverman W. (2007) The systematics of a small spineless Desmodesmus species, D. costato‐granulatus (Sphaeropleales, Chlorophyceae), based on its rDNA sequence analyses and cell wall morphology. Journal of Phycology 43: 378‐396. . [BIOLOGY section]
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8.1.11 Chlorella sp.
Figure 27‐ Chlorella emersonii, left picture control culture, right oicture N‐starved culture D. Reinecke, BGU
SYMBOLS: B, D, E, H, PIV
TAXONOMY
Phylum Class Order Family Genus Species
Chlorophyta Trebouxiophyceae Chlorellales Chlorellaceae Chlorella
Related Species Numbers of names and species: There are 71 species (and infraspecific) names in the database at present, of which 32 have been flagged as currently accepted taxonomically. C. acuminata, C. angustoellipsoidea, C. anitrata, C. anitrata var. minor, C. antartica, C. aureoviridis, C. autotrophica, C. botryoides, C. candida, C. capsulata, C. communis, C. conductrix, C. conglomerata, C. desiccata, C. ellipsoidea var. minor, C. ellipsoidea, C. emersonii var. rubescens, C. emersonii var. globosa, C. emersonii, C. faginea, C. fusca var. rubescens, C. fusca var. vacuolata, C. fusca, C. glucotropha, C. homosphaera, C. kessleri, C. kolkwitzii, C. lobophora, C. luteo‐viridis var. lutescens, C. luteo‐viridis, C. marina, C. miniata, C. minor, C. minutissima, C. mirabilis, C. mucosa, C. mutabilis, C. nocturna, C. oocystoides, C. ovalis, C. parasitica, C. parva, C. peruviana, C. protothecoides, C. protothecoides var. mannophila, C. pyrenoidosa var. tumidus, C. pyrenoidosa, C. pyrenoidosa var. duplex, C. regularis var. minima, C. reisiglii, C. reniformis, C. rugosa, C. saccharophila, C. saccharophila var. ellipsoidea, C. salina, C. sorokiniana, C. spaerckii, C. sphaerica, C. stigmatophora, C. subsphaerica, C. terricola, C. trebouxioides, C. variabilis, C. vulgaris f. minuscula, C. vulgaris f. suboblonga, C. vulgaris f. globosa, C. vulgaris var. viridis, C. vulgaris var. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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autotrophica, C. vulgaris var. tertia, C. vulgaris, C. zofingiensis.
BIOLOGY Structural and morphological features Chlorella is a large and diverse genus of small unicellular green algae of highest relevance to multiple aspects of biotechnology. Cells are round or ellipsoidal in shape. Smooth rigid cellulosic cell wall contains contains glucosamine (chitosan). Nucleus is single and eccentric, chloroplast single and parietal, pyrenoid single and covered with starch envelope. Pyrenoid stroma penetrated with 2 or 3 closely appressed thylakoids. Only asexual reproduction by autospores is known, autospores (2‐16 per mother cell) are released by rupture of parental cell wall. Chlorella, essentially cosmopolitan, occurs in both freshwater and marine habitats. Traditionally, the genus Chlorella has been comprised over 100 species; nevertheless, 10 well established species were recognized by chemotaxonomic methods (Kessler and Huus, 1992). Based on modern polyphasic approach, only four: C. vulgaris Beijerinck, C. lobophora Andreyeva, C. sorokiniana Shihira & Krauss, and C. kessleri Fott & Nováková (Huus et al., 1999) or even three: C. vulgaris, C. lobophora and C. sorokiniana (Krienitz et al., 2004) “true” Chlorella species are recognized recently. Strains of speciel interest Chlorella are fast growing freshwater, in some cases marine, water algae, reported to accumulate high concentration of oil under stress (Demirbas, 2009). Furthermore Chlorella strains can express a variety of carotenoids, among them astaxanthin. Some species is characterized by a very high growth rate (µmax = 0.20/h) and tolerance to a high culture temperature (40oC). Its desirable technological properties (resistance to shear stress, low adhesion to a surface of the bioreactor, low tendency to form aggregates) are expected to offer significant advantages for its use in large‐scale production bioreactors (Doucha and Lívanský, 2009). Its ability to grows under high CO2 permits with a direct supply of flue gas containing up to 40% (v/v) of CO2 (Doucha and Lívanský, 2005; Douskova et al., 2009, 2010). Chlorella vulgaris is a fast growing species and some strains can accumulate very high concentrations of lipids under stress (Francisco et al., 2010; Hsieh and Wu, 2009; Ly et al., 2010; Liang et al., 2009) while another othersones accumulate high amount of starch (Doušková et al., 2010). Oil production was tested also in other Chlorella species and strains as termophilic alga Chlorella sorokiana, or heterotrophically grown Chlorella protothecoides (Xu et al., 2006). Chlorella zofingiensis, can accumulate astaxanthin and lutein (Del Campo et al., 2004; Liu et al., 2010a,b; Yp et al., 2004). Biochemical composition and biomass main constituents The biochemical composition and the essential aminoacid content of Chlorella biomass are repoted in Tables 12 and 13. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Table 12 ‐ Basic chemical biomass composition of a production strain Chlorella sp, strain P 12. Item % algal DW Moisture 7‐7 Proteins (N x 6.25) 55‐58 *Lipids 8‐12 Saccharides 10‐15 Fibre 6‐8 Mineral substances 6‐8 Chlorophyll 2.5‐3.5 Nucleic acids 3‐4 *The proportion of essential unsaturated fatty acids (oleic, linoleic, linolenic) in the total fatty acids under optimum growth conditions is in the range of 40‐60 %.
Table 13 ‐ The percent proportion of essential amino acids in Chlorella and other protein‐rich sources. Amino acid Chlorella (dry weight) Isoleucine Leucine Lysine Methionine Phenylalanine Threonine Tryptophan Valine
2.01 4.14 3.19 1.04 2.57 2.42 0.80 3.00
In addition to 2‐3% of chlorophyll, Chlorella contains also carotenoids, orange and yellow dyespigments. The most valuable of these is beta‐carotene, provitamin A. The amount of β‐carotene in Chlorella is in the range of 0.10 to 0.25 % dry weight An important component of the Chlorella cells is biologically complexed, and therefore readily utilisedutilisable, basic minerals (phosphorus, potassium, magnesium, calcium and iron) (Table 14) and trace elements, which form part of enzyme complexes and vitamins. These elements include in particular manganese, zinc, molybdenum, copper and cobalt. Trace elements are often chelated with amino acids. Their concentration and type of binding can be considerably modified. This offers us the possibility to obtain algal biomass with a defined, mostly increased content of the desirable elements or their mixture in a natural organic form, which enhances their biological efficiency. Another group of substances which is present in Chlorella in much higher levels than in other plants is vitamins (Table 15). Striking is the high content of vitamins of group B, ascorbic acid (vitamin C), nicotinic acid (vitamin B3) and tocopherols (vitamin E).
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Table 14 ‐ Proportion of mineral substances and important trace elements in the Chlorella dry weight. Element mg/100 g DW Phosphorus 1200 Potassium 879 Sulphur 600 Magnesium 300 Calcium 230 Iron 70 Manganese 14 Zinc 11 Copper 4 Cobalt 0.5 Table 15 ‐ Vitamin content of Chlorella. Vitamins (mg kg‐1)
Chlorella (dry weight)
B1 ‐ thiamine B2 ‐ riboflavin B3 ‐ nicotinic acid (niacin) B5 ‐ pantothenic acid B6 ‐ pyridoxine B12 ‐ cobalamin biotin (vitamin H) folic acid vitamin E (tocopherol) vitamin C (ascorbic acid) β‐carotene (provitamin A)
18 44 219 13 28 0.8 0.3 42 298 655 1050
Chlorella Growth Factor (CGF) is a water‐extractable cell fraction containing free amino acids, peptides, glycoproteins, polyamines, some vitamins, minerals and other, as yet not exactly defined components. The effects of the extract are presented as striking, though scientific data to prove these statements are still at laboratory scale. It It promotes tissue regeneration, cell growth and division. It stimulates the production of leukocytes and their phagocytic activity, i.e. the ability to eliminate foreign bacteria and also the production of lymphocytes responsible for the synthesis of antibodies ‐ important factors in the immunity against infections. It is a suitable dietary supplement during the administration of probiotics, i.e. substances positively affecting the composition of intestinal microflora. It has been shown that, following an administration of the algal extract, the organism exhibits a better regeneration of damage caused by ionising radiation. Chlorella extracts have found their use in topical applications, e.g. in the treatment of chronic inflammations, eczemas, crural ulcers, burns and other badly healing wounds, which are healed by a fully functional tissue. Japanese laboratories have repeatedly published data on the anti‐tumour activity of the algal extract in vitro. The nutrient solution, in which Chlorella has been cultured, displays also a conspicuous stimulatory effect when used for watering freshly planted fruit or forest trees or vegetables. This has been attributed to its stimulatory effects on plant root‐taking and growth. The stimulatory effect is ascribed, apart from other components, to compounds of the phytohormone group which have been identified in the algal extracts. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Gross composition under optimal and stressed conditions A complex treatment of agricultural waste including the following major steps: anaerobic fermentation of suitable waste, cogeneration of the obtained biogas and growth of microalgae consuming the CO2 from biogas or flue gas, was verified under field conditions in a pilot‐scale photobioreactor. Laboratory analyses of the produced microalgae confirmed that it meets the strict EU criteria for relevant contaminants level in foodstuff (Doušková et al., 2009; 2010a; Kaštánek et al., 2010). The freshwater alga Chlorella, a highly productive source of starch, might substitute for starch‐rich terrestrial plants in bioethanol production. Cheap enhanced starch biomass can be produced from highly productive Chlorella cultures grown in suitable outdoor photobioreactors in which the photosynthetic carbon dioxide source is derived from combustion of organic waste, fermentation processes or other sources (Doucha et al., 2005; Douskova et al., 2009; Mann et al., 2009). This characteristic enhances the ecological and economic impact of the proposed technology, because of its potential to bioremediate carbon dioxide emissions from different CO2 sources including waste incinerators, power stations, limekilns, cogeneration units, etc. in situ.
BIOTECHNOLOGY Culture media Chlorella was one of the first algae isolated as a pure culture by Beijerinck in 1890. Since the half of the last century, attention has been drawn towards its potential for autotrophic mass cultivation. To this purpose, many mineral media were developed based in principle on the chemical composition of the algal cells. The basic inorganic elements used are: N, P, K, Mg, Ca, S, Fe, Cu, Mn, Mo and Zn (Krauss, 1958; O´Kelly, 1968). Many formulas are used for the cultivation of this genus ‐ see Hama and Miyachi (1988). For high‐yielded production of Chlorella outdoors an optimized composition of nutrient solution was proposed by Doucha and Lívanský (2006). The mineral medium was based on the mean content of P, N, K, Mg, and S in algal biomass and had the following initial composition (mg/L): 1100 (NH2)2CO, 237 KH2PO4, 204 MgSO4.7H2O, 40 C10H12O8N2NaFe, 88 CaCl2, 0.83 H3BO3, 0.95 CuSO4.5H2O, 3.3 MnCl2.4H2O, 0.17 (NH4)6Mo7O24.4H2O, 2.7 ZnSO4.7H2O, 0.6 CoSO4.7H2O, 0.014 NH4VO3 in distilled water. Cultivation of Chlorella heterotrophically in fermenters is also used (Lee et al., 1997). Heterotrophic culture may provide a cost effective, large‐scale alternative method for cultivation of some microalgae that can utilize organic carbon substances as their sole carbon and energy source (Chen and Chen, 2006). Nutrient solution containing glucose or acetate as a source of carbon for intensive growth of Chlorella in fermenter was described by Endo and Shirota (1972) and lately by Doucha and Lívanský (Cz. patent. 288638, 2001). Technology of heterotrophic Chlorella cultivation has been commercially used for production of biomass (Doucha and Lívanský, 2011) for food and feed as well as for production of biomass enriched by organically bound selenium (Doucha et al., 2009;, Skřivan et al., 2006; Trávníček et al., 2008) or as a source of lipids for production of biodiesel (Xiong et al., 2008). Cultivation methods and production systems There is still a noticeable discrepancy between the extent of commercially operated algal cultures and the potential of algae. Since the first experiments with large‐scale algal cultures in the 1950s (Burlew, 1953) many types of culture equipment have been developed (Stengel, 1970; Richmond and Becker, 1986; Tredici, 2004). For most products of microalgal mass cultivation outdoor open circular or “raceway” ponds with a 15‐30 cm layer of algal suspension are the most used technology for the growth of algae AquaFUELs‐ Taxonomy, Biology and Biotechnology
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(Borowitzka, 1999). The commercial production of Chlorella biomass is carried out exclusively in these open systems. Closed reactors (tubular, helical and flat bioreactors, vertical cylinders and sleeves) have been employed for research in small field installations (e.g. Torzillo, 1997; Pulz and Scheibenbogen, 1998; Tredici, 2004). The only exception is a large‐scale tubular bioreactor which started production of Chlorella biomass in Central Germany in 2000 (Pulz, 2001) and tubular reactors used to produce Haematococcus in Israel. Open ponds are characterized by simple construction and relatively low building costs. On the other hand, there are many serious drawbacks of this system: (a) due to thick layer of algal suspension, the culture must be grown at low densities (about 0.5 algal DW l‐1). With increasing densities the productivity sharply decreases; (b) low velocity flow (15‐30 cm s‐1) of poorly mixed algal suspension leads to low utilization of light energy and to accumulation of oxygen dissolved in the suspension; (c) the separation of low concentrated algae from the nutrient solution at the harvest is highly energy demandinged is the separation of low concentrated algae from the nutrient solution at the harvest. The key for reduction of cultivation costs rests in a low area volume of algal suspension. This can be achieved by decreasing algal layer to a low value as technologically possible (Doucha and Lívanský, 2006, 2009). Technological and production characteristics of both systems are given below (Table 16). Table 16‐ Characteristics of cultures in raceway ponds and thin layer culture system. Culture characteristics culture volume (l m‐2)
Raceway ponds 150‐300
Thin‐layer 6‐8
culture layer thickness (mm) biomass harvest density (g l‐1)
150‐300 0.5‐1
6‐8 35‐50
harvest/downstream processing density multiply surface/volume ratio (m‐1)
150‐300
3‐4.3
5
100
2.5‐4
5‐8
10‐20
20‐40
0.05‐0.1 5‐20
2‐5 60‐70
photosynthetic efficiency (% of PAR) areal productivity (g m‐2 d‐1) volumetric productivity (g l‐1 d‐1) efficiency of CO2 utilization (%)
Harvesting methods Many methods are available for harvesting of microalgae, consisting in thickening of algal biomass as a first step. These include: centrifugation, electroflotation, and chemical flocculation, followed by sedimentation or air flotation, continuous belt filtration, vibrating and stationary screens, sand bed filtration, and autoflocculation (Richmond, 1986). Speaking about large‐scale commercial Chlorella cultures, only centrifugation by means of continuously operating self cleaning centrifuges, is used. The advantage of centrifugation is its simplicity and possibility to lower chemicals and bacterial contamination in product. On the other side, this process is connected with a high energy demand. When commonly used raceway pond culture technology is used (at harvesting density about 0.5 g l‐1 and thickening up to 150 g l‐1), about 30 % of the total cost of the production is accounted (Gudin and Thepenier, 1986). Biomass processing The next step after the thickening of algal biomass by centrifugation is disruption of algal cells. The rigid cellulosic cell wall, one of the characteristics of unicellular Chlorella, causes a low utilization of cell content by a recipient. The digestibility of ruptured Chlorella cells increases to 80 % (Doucha and Lívanský, 2008) AquaFUELs‐ Taxonomy, Biology and Biotechnology
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compared to 15‐25 % for unruptured cells (Becker, 1984). To open the cell, methods of freezing, alcalic alkaline and organic solvents, osmotic shocks, sonication, high‐pressure homogenization and bead milling were tested (Molina Grima et al., 2004). For large‐scale processing of Chlorella cultures disintegration of cell walls by bead mills is mostly used (Middelberg, 1995; Doucha and Lívanský, 2008). Spray drying as the end step of the downstream processing process is the most extended method for dehydration of ruptured Chlorella cells. Algae biomass is dispersed into very fine droplets, whose surface temperature in the course of several seconds of drying does not surpass 60 oC. Thus, the process is very considerate and the quality of the product is high. Scaling up The first commercial production of Chlorella and, some years later, blue‐green Spirulina cultures started in Japan and Taiwan in the 1960’s. Nowadays, large‐scale production plants are located also in the USA, China, India, Thailand, Indonesia, Germany and other countries (Borowitzka, 1996; Lee, 1997; Pulz , 2001; Borowitzka 1996; Tredici, 2004; Spolaore et al., 2006). Their per year ‐ world production is estimated to be about 8000 tons (Spirulina) and 5000 tons (Chlorella). Most of the products have been used in human nutrition. High growth rate, high photosynthetic efficiency, relatively high content of energy‐rich chemicals on one side and experience with large‐scale culture and downstream processing technologies concentrate in last decades increasing attention on microalgae as a feedstock for biofuels. Today projects dealing with algae are focused almost entirely on biodiesel production. Nevertheless, algal strains containing higher amount of lipids are characterized by low growth rate. Slow growth increases the operational costs and demands cultivation in closed bioreactors whose building is expensive. On the other side for economical production of bioethanol, relatively cheap biomass of high‐yielded Chlorella cultures, containing enhanced amount of starch grown in suitable open bioreactors is perspective solution. To produce starch economically, conditions for culturing starch‐enriched algae in dense cultures must be attained. Using a thin layer algal suspension in outdoor cultures, linear growth continues up to very high biomass concentrations (about 40 g/L) enabling easy and cheap harvesting and processing (Doucha and Lívanský, 2006, 2009). However, the content of starch in the biomass is low (15% of DW or less). The conditions under which starch content increases in commercially produced algal biomass to a level that would be viable for bioethanol production can be achieved if the processes and events during which starch is extensively degraded are slowed down, or stopped completely, while the factors supporting starch synthesis, namely light intensity, remain sustainable (Figure 27) (Brányiková et al., 2011). Similarly, the Chlorella species producing oil as their energy reserves rather than starch can be similarly treated to markedly increase their oil content (Figure 28). An increase in the production of starch in sulphur‐limited culture up to a maximum of 50% starch content of algal biomass (DW) was demonstrated under field conditions using the outdoor scale up, thin‐ layer solar photobioreactor. While use of algae with enriched starch content is conventional for bioethanol production, another attractive exploitation of starch from algae might be the production of hydrogen, which may be realized in the near future (Miura et al., 1982; Tsygankov et al., 2002; Melis et al., 2004; Chochois et al., 2009; Melis et al. 2004; Miura et al. 1982; Tsygankov et al. 2002). Sulphur limitation could be one of the ways to support hydrogen production (Melis et al., 2000; Zhang et al., 2002). It has been shown recently that some strains of Chlorella can produce and accumulate significant volume of hydrogen gas under anaerobic conditions and sulphur deprivation such as it is reported in literature using C. reinhardtii (Chader et al., 2009). AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Figure 28 ‐ Electron microscopic photographs of daughter (A) and mother (B) cells of Chlorella grown in complete mineral medium, in the presence of cycloheximide (1 mg/L) (C), and in sulfur limiting medium (D). N nucleus, S starch granules. Bars: panels A, B, C = 2 μm; bar panel D = 5 μm.
Figure 29 ‐ Fluorescent microphotography of Chlorella cells with enriched content of oil (yellow spots oil stained by Nile Red
To decrease price of algal biomass the flue gas from various sources can be used as a cheap CO2 source. Using pilot outdoor thin‐layer bioreactor, built in a livestock farm, flue gas, after utilization of CH4 anaerobically generated in biogas station, was used as a source of CO2 for algal photosynthesis. Besides, minerals of liquid concentrate of the anaerobic digested livestock excrements can be used as a source of inorganic nutrients for algal growth (Doucha, 2010, unpublished results). The flue gas, after utilization of biogas produced from distillery stillage (Doušková et al., 2010) or from swine manure (Kaštánek et al., 2010) for electricity and heat production, was also successfully applied as a cheap source of CO2 for algal biomass production. An increase in the production of starch in sulfur‐limited culture up to a maximum of 50% starch content of algal biomass (DW) was demonstrated under field conditions using the outdoor scale up, thin‐layer solar photobioreactor. Despite the relatively unfavourable climatic conditions of Trebon (Czech Republic), a total yield of starch calculated per ha over a season of 150 days was 7 tonnes (Doušková et al., 2010). In optimum locations for photoautotrophic production of algae like Greece, with a season lasting approximately 250 days, the overall harvest might be increased by a factor of 10 (Doucha and Livansky, 2006). The remaining parts of the cells, containing largely proteins, can be used as a feed supplement what further decreases the cost of starch production. References Bayen M., Dalmon J. (1975) [Physico‐chemical determination of the ploidy of the unicellular alga, Chlorella pyrenoidosa (strain 211/8b) (author's transl)]. Biochimica et Biophysica Acta 395: 213‐219. Becker E.W (1994) Microalgae: Biotechnology and Microbiology. Cambridge University Press, Cambridge. Borowitzka M.A. (1996) Closed algal photobioreactors: Design considerations for large‐scale systems. Journal of Marine Biotechnology 4: 185‐191.
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Borowitzka M.A. (1999) Commercial production of microalgae: ponds, tanks, tubes and fermenters. Journal of Biotechnology 70: 313‐321. Brányiková I., Maršálková B., Doucha J., Brányik T., Bišová K., Zachleder V., Vítová V. (2011) Microalgae ‐ novel highly‐efficient starch producers. Biotechnology and Bioengineering 108: 766‐776. Burlew J.S. (ed.) (1953) Algal Culture from Laboratory to Pilot Plant. Carnegie Institute of Washington, DC. Chader S., Hacene H., Agathos S.N. (2009) Study of hydrogen production by three strains of Chlorella isolated from the soil in the Algerian Sahara. International Journal of Hydrogen Energy 34: 4941‐4946. Chen G.Q., Chen F. (2006) Growing phototrophic cells without light. Biotechnology Letters 28: 607‐616. Chochois V., Dauvillée D., Beyly A., Tolleter D., Cuiné S., Timpano H., Ball S., Cournac L., Peltier G. (2009) Hydrogen production in Chlamydomonas: photosystem II‐dependent and independent pathways differ in their requirement for starch metabolism. Plant Physiology 151: 631‐640 Del Campo J.A., Rodriguez H., Moreno J., Vargas M.A., Rivas J., Guerrero M.G. (2004) Accumulation of astaxanthin and lutein in Chlorella zofingiensis (Chlorophyta). Applied Microbiology and Biotechnology 64: 848‐854. Demirbas A. (2009) Production of biodiesel from algae oils. Energy Sources, Part A: Recovery, utilization, and environmental effects 31: 163‐168. Doucha J., Lívanský K. (2001) Method of controlled cultivation of algae in heterotrophic mode of nutrition. Czech Patent 288638. Doucha J., Lívanský K. (2006) Productivity, CO2/O2 exchange and hydraulics in outdoor open high density microalgal (Chlorella sp.) photobioreactors operated in a Middle and Southern European climate. Journal of Applied Phycology 18: 811‐826. Doucha J., Lívanský K. (2009) Outdoor open thin‐layer microalgal photobioreactor: potential productivity. Journal of Applied Phycology 21: 111‐117. Doucha J., Lívanský K. (2008) Influence of processing parameters on disintegration of Chlorella cells in various types of homogenizers. Applied Microbiology and Biotechnology 81: 131‐440. Doucha J, Lívanský K. (2011) Production of high‐density Chlorella culture grown in fermenters. Journal of Applied Phycology DOI 10.1007/s10811‐010‐9643‐2. Doucha J., Straka F., Livansky K. (2005) Utilization of flue gas for cultivation of microalgae (Chlorella sp.) in an outdoor open thin‐ layer photobioreactor. Journal of Applied Phycology 17: 403‐412. Doucha J., Lívanský K., Kotrbáček V., Zachleder V. (2009) Production of Chlorella biomass enriched by selenium and its use in animal nutrition: a review. Applied Microbiology and Biotechnology 83: 1001‐1008. Doušková 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. Applied Microbiology and Biotechnology 82: 179‐185. Doušková I., Kaštánek F., Maléterová Y., Kaštánek P., Doucha J., Zachleder V. (2010) Utilization of distillery stillage for energy generation and concurrent production of valuable microalgal biomass in the sequence: Biogas‐cogeneration‐microalgae‐ products. Energy Conversion and Management 51: 606‐611. Endo H., Shirota M. (1972) Studies on the heterotrophic growth of Chlorella in a mass culture. Proceedings IV IFS: Fermentation Technology Today, pp. 533‐541. Francisco É.C., Neves D.B., Jacob‐Lopes E., Franco T.T. (2010) Microalgae as feedstock for biodiesel production: Carbon dioxide sequestration, lipid production and biofuel quality. Journal of Chemical Technology and Biotechnology 85: 395‐403. Guckert J.B., Cooksey K.E. (1990) Triglyceride accumulation and fatty acid profile changes in Chlorella (Chlorophyta) during high pH‐ induced cell cycle inhibition. Journal of Phycology 26: 72‐79. Gudin C., Thepenier C. (1986) Bioconversion of solar energy into organic chemicals by microalgae. Advances in Biotechnology Processes 6: 73‐110. Hsieh C.H., Wu W.T. (2009) Cultivation of microalgae for oil production with a cultivation strategy of urea limitation. Bioresource Technology 100: 3921‐3926. Huss V.A.R., Frank C., Hartmann E.C., Hirmer M., Kloboucek A., Seidel B.M., Wenzeler P., Kessler E. (1999) Biochemical taxonomy and molecular phylogeny of the genus Chlorella sensu lato (Chlorophyta). Journal of Phycology 35: 587‐598. Kaštánek F., Šabata S., Šolcová O., Maléterová .Y, Kaštánek .P, Brányiková I., Kuthan K., Zachleder V. (2010) In‐field experimental verificatiom of cultuvation of microalgae Chlorella sp. using the flue gas from a cogeneration unit as a source of carbon dioxide. Waste Management and Research 28: 961‐966. Kessler E., Huss V.A.R. (1992) Comparative physiology and biochemistry and taxonomic assignment of the Chlorella (Chlorophyceae) strains of the Culture Collection of the University of Texas at Austin. Journal of Phycology 28: 550‐553. Krauss R.W. (1958) Physiology of the fresh‐water algae. Annual Review of Plant Physiology 9: 207‐244. Krienitz L.., Hegewald E.H., Hepperle D., Huss V.A.R., Rohr T., Wolf M. (2004) Phylogenetic relationship of Chlorella and Parachlorella gen. nov. (Chlorophyta, Trebouxiophyceae). Phycologia 43: 529‐542. Krusteva N.G., Tomova N.G., Georgieva M.A. (1984) Allosteric regulation of NAD(NADP)‐dependent glyceraldehyde‐3‐phosphate dehydrogenase from Chlorella by α‐amino acids, dithiothreitol and ATP. FEBS Letters 171: 137‐140. Lee Y.K. (1997) Commercial production of microalgae in the Asia‐Pacific rim. Journal of Applied Phycology 9: 403‐411. Lee Y.K. (2001) Microalgal mass culture systems and methods: Their limitation and potential. Journal of Applied Phycology 13: 307‐ 315. Li X., Xu H., Wu Q. (2007) Large‐scale biodiesel production from microalga Chlorella protothecoides through heterotrophic cultivation in bioreactors. Biotechnology and Bioengineering 98: 764‐771. Liang Y., Sarkany N., Cui Y. (2009) Biomass and lipid productivities of Chlorella vulgaris under autotrophic, heterotrophic and mixotrophic growth conditions. Biotechnology Letters 31: 1043‐1049. Liu J., Huang J., Fan K.W., Jiang Y., Zhong Y., Sun Z., Chen F. (2010a) Production potential of Chlorella zofingienesis as a feedstock for biodiesel. Bioresource Technology 101: 8658‐8663.
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Liu J., Zhong Y., Sun Z., Huang J., Sandmann G., Chen F. (2010b) One amino acid substitution in phytoene desaturase makes Chlorella zofingiensis resistant to norflurazon and enhances the biosynthesis of astaxanthin. Planta 232: 61‐67. Mann G., Schlegel M., Schumann R., Sakalauskas A. (2009) Biogas‐conditioning with microalgae. Agronomy Research 7: 33‐38. Melis A., Seibert M., Happe T. (2004) Genomics of green algal hydrogen research. Photosynthesis Research 82: 277‐288. Melis A., Zhang L., Forestier M., Ghirardi M.L., Seibert M. (2000) Sustained photobiological hydrogen gas production upon reversible inactivation of oxygen evolution in the green alga Chlamydomonas reinhardtii. Plant Physiology 122: 127‐135. Middelberg A.P.J. (1995) Process‐scale disrruption of microorganisms. Biotechnology Advances 13: 491‐555. Miura Y., Yagi K., Shoga M., Miyamoto K. (1982) Hydrogen production by a green alga, Chlamydomonas reinhardtii, in an alternating light/dark cycle. Biotechnology and Bioengineering 24: 1555‐1563. Molina Grima E., Fernández F.G.A., Medina A.R. (2004) Downstream processing of cell‐mass and products. In: Richmond A. (ed.) Handbook of Microalgal Culture: Biotechnology and Applied Phycology. Blackwell Science, pp 215‐251. O´Kelly J.C. (1968) Mineral nutrition of algae. Annual Review of Plant Physiology 19: 89‐112. Oh‐hama T., Miyachi S. (1988) Chlorella. In: Borowitzka M.A., Borowitzka L.J. (eds.), Micro‐algal Biotechnology. Cambridge University Press, Cambridge, pp. 3‐26. Pulz O. (2001) Photobioreactors: Production systems for phototrophic microorganisms. Applied Microbiology and Biotechnology 57: 287‐293. Pulz O., Scheibenbogen K. (1998) Photobioreactors: Design and performance with respect to light energy input. In: Scheper T. (ed.) Advances in Biochemical Engineering/Biotechnology, Springer‐Verlag, Berlin, pp. 123‐152. Richmond A. (ed.) (1986) CRC Handbook of Microalgal Mass Culture. CRC Prsss, Boca Raton, Florida. Richmond A., Becker E.W. (1986) Technological aspects of mass cultivation – a general outlook. In: Richmond A. (ed.) CRC Handbook of Microalgal Mass Culture. CRC Press, Boca Raton, Florida, pp 245‐263. Rodolfi L., Chini Zitteli G., Bassi N., Padovani G., Biondi N., Bonini G., Tredici M.R. (2009) Microalgae for oil: Strain selection, induction of lipid synthesis and outdoor mass cultivation in a low‐cost photobioreactor. Biotechnology and Bioengineering 102: 100‐112. Sauer N., Tanner W. (1989) The hexose carrier from Chlorella. cDNA cloning of a eukaryotic H+‐cotransporter. FEBS Letters 259: 43. Skřivan M., Šimáně J., Dlouhá G., Doucha J. (2006) Effect of dietary sodium selenite, Se‐enriched yeast and Se‐enriched Chlorella on egg Se concentration, physical parameters of eggs and laying hen production. Czech Journal of Animal Science 51: 163‐167. Spolaore P., Joannis‐Cassan C., Duran E., Isambert A. (2006) Commercial application of microalgae. Journal of Bioscience and Bioengineering 101: 87‐96. Stengel E. (1970) Anlagentype und Verfahren der technischen Algenmassenproduktion. Berichte der Deutschen Botanische Gesellschaft 83: 589‐606. Syrett P.J., Thomas E.M. (1973) The assay of nitrate reductase in whole cells of Chlorella: Strain differences and the effect of cell walls. New Phytologist 72: 1307‐1310. Torzillo G. (1997) Tubular bioreactors. In: Vonshak A (ed.) Spirulina platensis (Arthrospira): Physiology, Cell Biology and Biotechnology. Taylor & Francis, London, pp 101‐115. Trávníček J., Racek J., Trefil L., Rodinová H., Kroupová V., Illek J., Doucha J., Písek L. (2008) Activity of glutathione peroxidase (GSH‐ Px) in the blood of ewes and their lambs receiving the selenium‐enriched unicellular alga Chlorella. Czech Journal of Animal Science 53: 292–298. Tredici M.R. (2004) Mass Production of Microalgae: Photobioreactors. In: Richmond A. (ed.) Handbook of Microalgal Culture. Blackwell Science Ltd, Oxford, pp 178‐214. Tsygankov A., Kosourov S., Seibert M., Ghirardi M.L. (2002) Hydrogen photoproduction under continuous illumination by sulfur‐ deprived, synchronous Chlamydomonas reinhardtii cultures. International Journal of Hydrogen Energy 27: 1239‐1244. Xiong W., Li X., Xiang F., Wu Q. (2008) High density fermentation of micro alga Chlorella protothecoides in bioreactor for microbio‐ diesel production. Applied Microbiology and Biotechnology 78: 29‐36. Xu H., Miao X., Wu Q. (2006) High quality biodiesel production from a microalga Chlorella protothecoides by heterotrophic growth in fermenters. Journal of Biotechnology 126: 499‐507. Yp J.M., Cheng L.H., Xu X.H., Zhang L., Chen H.L. (2010) Enhanced lipid production of Chlorella vulgaris by adjustment of cultivation conditions. Bioresource Technology 101: 6797‐6804. Yp P.F., Wong K.H., Chen F. (2004) Enhanced production of astaxanthin by the green microalga Chlorella zofingiensis in mixotrophic culture. Process Biochemistry 39: 1761‐1766. Zhang L., Happe T., Melis A. (2002) Biochemical and morphological characterization of sulfur‐deprived and H2‐producing Chlamydomonas reinhardtii (green alga). Planta 214: 552‐561.
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8.1.12 Parietochloris incisa
Figure 30 – Parietochloris incisa in balanced growth (left) and nitrogen starved (right) I. Khozin, BGU
SYMBOLS: D, PIV
TAXONOMY
Phylum Class Order Family Genus Species
Chlorophyta Trebouxiophyceae Chlorellales Chlorellaceae Parietochloris Parietochloris incisa
Related Species Numbers of names and species: There are 6 species (and infraspecific) names in the database at present. P. alveolaris, P. bilobata, P. cohaerens, P. incisa, P. ovoidea, P. pseudoalveolaris.
BIOLOGY Parietochloris incisa is a unicellular, oleaginous, freshwater alga. The alga was isolated from the slopes of a snow mountain in Japan (Watanabe et al., 1996), an alpine environment which is characterized by a broad temperature range, UV radiation and light levels that can be extremely high. Such environments were found to be a natural habitat for phototrophic microorganisms that accumulate high polyunsaturated fatty acids (PUFA) concentrations, including sea ice diatoms, dinoflagellates, and green algae. Indeed P. incisa was found to accumulate unusually high content of the long chain PUFA, arachidonic acid (AA) (Bigogno et al., 2002a).
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Biochemical composition Under nitrogen starvation, the fatty acid content in P. incisa is over 35% of dry weight; AA constitutes about 60% of total fatty acids, and over 90% of cell AA is deposited in TAG (Khozin‐Goldberg et al., 2002). In the lipid‐linked pathway of its biosynthesis, AA is mainly exported to TAG and accumulated in cytoplasmic lipid bodies (Bigogno et al., 2002b). The pathway of AA biosynthesis involves stepwise desaturation and elongation of oleic acid via the so‐called ω6 pathway. The genes encoding for ∆12, ∆6, ∆5 desaturases and ∆6 PUFA elongase were cloned and their functions were validated in heterologous system (Iskandarov et al., 2009, 2010). Furthermore, under optimal growth temperature (25 °C), AA is partially converted to the valuable PUFA eicosapentaenoic acid (EPA, 20:5ω3). This conversion is more pronounced when alga is exposed to low temperature (10‐15oC). The conversion of ω6 precursors into ω3 fatty acids is catalyzed by a group of enzymes called ω3 fatty acid desaturases (FAD) that differ by fatty acid substrate preference and cellular localization. A Δ5‐desaturase‐deficient mutant isolated at BGU (Cohen et al., 2009) is able to accumulate the high value PUFA DGLA (20:3ω6).
BIOTECHNOLOGY Growth medium mBG11 Growth rate Growth rate in panel reactors and continuous illumination under nitrogen starvation: 160 mg/l per day accumulating 6% DGLA and 18% TAG. Pilot scale scale production The DGLA production process using the P. incisa mutant has been tested at BGU at the pilot scale using 50 – 200 l flatpanel reactors in greenhouse or climate controlled growth rooms. DGLA content 10% of total dry biomass was achieved after cultivation in N‐deficient medium demonstrating a commercial production capability. HIGHLIGHTS IN BIOTECHNOLOGY P. incisa and its mutant are unique among algae in being able to accumulate high concentrations of PUFA in TAG in cytoplasmic oil globules. Identification of several desaturase and elongase genes may allow breakthroughs in engineering more efficient PUFA producing algae. References Bigogno C., Khozin‐Goldberg I., Boussiba S., Vonshak A., Cohen Z. (2002a) Lipid and fatty acid composition of the green oleaginous alga Parietochloris incisa, the richest plant source of arachidonic acid. Phytochemistry 60: 497‐503. Bigogno C., Khozin‐Goldberg I., Adlerstein D., Cohen Z. (2002b) Biosynthesis of arachidonic acid in the oleaginous microalga Parietochloris incisa (Chlorophyceae): radiolabeling studies. Lipids 37: 209‐216. Cohen Z., Khozin‐Goldberg I., Boussiba S., Vonshak A.; Ben‐Gurion University of the Negev Research and Development Authority, assignee. 2009 19.02.2009. Over‐Production of dihomo gamma linolenic acid by a Mutant Strain of Parietochloris incisa. IL. Iskandarov U., Khozin‐Goldberg I., Ofir R., Cohen Z. (2009) Cloning and characterization of the ω6 polyunsaturated fatty acid elongase from the greenm Parietochloris incisa. Lipids 44: 545‐554. Watanabe S., Hirabayashi S., Boussiba S., Cohen Z., Vonshak A., Richmond A. (1996) Parietochloris incisa comb.nov. (Trebouxiophyceae, Chlorophyta). Phycological Research 44: 107‐108.
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8.1.13 Prototheca sp.
Figure 31 ‐ Light microscopic view showing Prototheca moriformis Lactophenol cotton blue mount fixation. Picture from http://content9.eol.org/content/2009/11/25/02/75871_small.jpg
SYMBOLS: D
TAXONOMY
Phylum Class Order Family Genus Species
Chlorophyta Trebouxiophyceae Chlorellales Chlorellaceae Prototheca
Related species There are 21 species (and infraspecific) names in the database at present, of which 13 have been flagged as currently accepted taxonomically. P. blaschkeae, P. chlorelloides, P. ciferrii, P. crieana, P. filamenta, P. hydrocarbonea, P. kruegeri, P. moriformis, P. moriformis var. betulina, P. portoricensis, P. portoricensis var. trisporus, P. salmonis, P. segbwema, P. stagnorum, P. trispora, P. ulmea, P. viscosa, P. wickerhamii, P. zopfii var. portoricensis, P. zopfii var. hydrocarbonea, P. zopfii.
BIOLOGY The genus Prototheca is composed of microscopic achlorophyllous organisms with a life cycle similar to that of the genus Chlorella. The species typically produce thick‐walled cells (sporangia) which, at ma‐turity, divide by irregular cleavage forming 2‐15 aplanospores (endo‐spores). Following rupture of the sporangial wall, freed aplanospores enlarge and repeat the cycle. One to three percent of the sporangia cleave to form 2‐3 thick‐walled resting cells (hypnospores). No sexual cycle has been observed. Prototheca can use dextrose, levulose, galactose, ethanol, n‐butanol, iso‐butanol, iso‐pentanol, hexanol and glycerol and some species assimilate sucrose, trehalose, n‐propanol and n‐pentanol. In general, alcohols are assimilated similarly in both liquid and vapor phases; n‐pentanol is assimilated only in the vapor phase. Species of Prototheca require thiamine. AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Numerous studies have reported a pathogenic potential for P. wickerhamii and P. zopfii. The cases of human protothecosis are predominantly caused by P. wickerhamii and occur as local (predominantly cutaneous) and systemic infections mainly in immune‐compromised patients, e.g. patients infected with HIV or treated with glucocorticoids. P. blaschkeae were isolated from some cases of onychomycosis. Canine protothecosis is caused by P. wickerhamii and P. zopfii, and is characterized by similar clinical symptoms as in humans. Worldwide, P. zopfii has been identified to induce a therapy‐resistant inflammation of the mammary gland in dairy cows.
BIOTECHNOLOGY P. zopfii and P. moriformis have been proposed, together with Chlorella protothecoides, as organisms for production of ascorbic acid (Running et al., 2002). In a laboratory scale experiment (1‐14 L fermentors) P. zopfii at 35 °C at pH 7 produced 37.8 mg L‐1 of ascorbic acid, mainly intracellular, with a biomass preodcution of 27 g L‐1, while at pH 4/5 it produced 73 mg L‐1 of extracellular ascorbic acid and 55 g L‐1 of biomass in 29 h; similarly P. moriformis produced in 30 h at pH 4 162 mg L‐1 of extrcellular ascorbic acid and 42 g L‐1 of biomass (Huss et al., 1995; Running, 1999). P. zopfii is able to utilise crude oil hydrocarbons (aromatics, cyclic and branched alkanes) as well as pure n‐alkanes (Ueno et al., 2002, 2008). Solazyme Inc. (San Francisco, USA) has recently patented several processes to obtain oil from microrganisms including Prototheca, that contains lipids with a higher degree of saturation compared to other algae and has the advantage of lacking pigments, also through genetic engineering to increase production (Dillon et al., 2010; Franklin et al., 2010, 2011). References Arnold P., Ahearn D.G. (1972) The systematics of the genus Prototheca with a description of a new species P. filamenta. Mycologia 64: 265‐275. [BIOLOGY section] Dillon H.F., Elefant D., Day A.G., Franklin S., Wittenberg J. (2010) Fractionation of oil‐bearing microbial biomass. WO2010/138620. Franklin S., Somanchi A., Espina K., Rudenko G., Chua P. (2011) Renewable chemical production from novel fatty acids feedstocks. US Patent No. 7,883,882 B2. Franklin S., Somanchi A., Espina K., Rudenko G., Chua P. (2010) Manufactoring of tailored oils in recombinant heterotrophic microorganisms. WO2010/063031 A2. Huss J.R., Running J.A., Skatrud T.J. (1995) L‐ascorbic acid production in microorganisms. WO95/21933. Running J.A. (1999) Process for the production of ascorbic acid with Prototheca. US Patent No. 5,900,370. Running J.A., Severson D.K., Schneider K.J. (2002) Extracellular production of L‐ascorbic acid by Chlorella protothecoides, Prototheca species, and mutants of P. moriformis during aerobic culturing at low pH. Journal of Industrial Microbiology and Biotechnology 29: 93‐98. Sudman M.S., Kaplan W. (1973) Identification of the Prototheca Species by immunofluorescence. Applied and Environmental Microbiology 25: 981‐990. [BIOLOGY section] Ueno R., Urano N., Wada S., Kimura S. (2002) Optimization of heterotrophic culture conditions for n‐alkane utilization and phylogenetic position based on the 18S rDNA sequence of a thermotolerant Prototheca zopfii strain. Journal of Bioscience and Bioengineering 94: 160‐165. Ueno R., Wada S., Urano N. (2008) Repeated batch cultivation of the hydrocarbon –degrading, micro‐algal strain Prototheca zopfii RND16 immobilised in polyurethane foam. Canadian Journal of Microbiology 54: 66‐70. von Bergen M., Eidner A., Schmidt F., Murugaiyan J., Wirth H., Binder H., Maier T., Roesler U. (2009) Identification of harmless and pathogenic algae of the genus Prototheca by MALDI‐MS. Proteomics Clinical Applications 3: 774‐784. [BIOLOGY section] Wolff G., Plante I., Lang B. F., Kück U., Burger G. (1994) Complete sequenceof the mitochondrial DNA of the chlorophyte alga Prototheca wickerhamii: Gene content and genome organization. Journal of Molecular Biology 237: 75‐86. [BIOLOGY section]
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8.2 Rhodophyta 8.2.1
Porphyridium cruentum
Figure 32 ‐ Light microscopic view showing Porphyridium cruentum strain CCALA 415. Picture from www.butbn.cas.cz/ccala/col_images/415.jpg
SYMBOLS: D
TAXONOMY
Phylum Class Order Family Genus Species
Rhodophyta Porphyridiophyceae Porphyridiales Porphyridiaceae Porphyridium Porphyridium cruentum
Related species There are 7 species (and infraspecific) names in the database at present, of which 3 have been flagged as currently accepted taxonomically. P. aerugineum, P. cruentum, P. griseum, P. marinum, P. purpureum, P. sordidum, P. violaceum.
BIOLOGY Porphyridium is composed of spherical to ovoid unicells with a stellate chloroplast and prominent central pyrenoid. The cell diameter is 5‐10 μm in the exponential phase, 7‐16 μm in the stationary phase. Cells are solitary, but often grouped into irregular colonies with an ill‐defined mucilaginous matrix. Species are distinguished by chloroplast color. The chloroplasts of freshwater species contain single thylakoids with phycobilisomes (granules consisting of the accessory pigments) on both sides. The phycobilisomes of blue‐ colored species, such as Porphyridium aerugineum, tend to be hemidiscoidal in shape and predominated by the blue pigment phycocyanin . In contrast, the phycobilisomes of the red‐colored Porphyridium purpureum AquaFUELs‐ Taxonomy, Biology and Biotechnology
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are larger and hemispherical and composed mostly of the red pigment phycoerythrin. Reproduction occurs by cell division. Porphyridium forms gelatinous coatings on various surfaces; it is widespread in freshwaters, brackish environments. Species of Porphyridium can form gelatinous crusts on moist soils and decaying wood . In these habitats, these species are reasonably desiccation resistant and shade tolerant.
BIOTECHNOLOGY The marine microalga Porphyridium cruentum is of increasing interest as source of valuable compounds like phycoerythrin, sulfated exopolysaccharides, superoxide‐dismutase, and polyunsaturated fatty acids with applications in the food, pharmaceutical and cosmetic industries (Fábregas et al., 1998; Dillon et al., 2007). Phycoerythrin is used as fluorescent dye in immunoassays (Bermejo Román et al., 2002), sulfated polysaccharides are known inhibit viruses (Huheihel et al., 2002) and show hypocholesterolemic activity in rats (Dvir et al., 2000, 2009). P. cruentum is also considered to be one of the most important sources of the polyunsaturated fatty acids eicosapentaenoic acid (20:5 ω 3, EPA) and arachidonic acid (20:4ω6, ARA) (Cohen and Cohen 1991; Guil‐Guerrero et al., 2001). In a 220‐L ailift tubular photobioreactor P. cruentum reached average biomass productivities of 1.76 g L‐ 1 day‐1 with 2% biomass dry weight of ARA , 1.7% palmitic acid and 1.1% EPA (Rebelloso Fuentes et al., 1999), similar to those obtained by Camacho Rubio et al. (1999). Porphyridium have been tested in screening for algal strains suitable for biodiesel production or considered as a potential interesting alga based on its biochemical composition (Griffiths and Harrison, 2009; Rodolfi et al., 2009), however up to now no in depth studies on this topic have been carried out. References Bermejo Román R., Alvárez‐Pez J.M., Acién Fernández F.G., Molina Grima E. (2002) Recovery of pure B‐phycoerythrin from the microalga Porphyridium cruentum. Journal of Biotechnology 93: 73–85. Camacho Rubio F., Acién Fernández F.G., Sánchez Pérez J.A., García Camacho F., Molina Grima E. (1999) Prediction of dissolved oxygen and carbon dioxide concentration profiles in tubular photobioreactors for microalgal culture. Biotechnology and Bioengineering 62: 71‐86. Cohen Z., Cohen S. (1991) Preparation of eicosapentaenoic acid (EPA) concentrate from Porphyridium cruentum. Journal of the American Oil Chemists' Society 68: 16‐19. Cohen Z., Vonshak A., Richmond A. (1988) Effect of environmental conditions on fatty acid composition of the red alga Porphyridium cruentum: correlation to growth rate. Journal of Phycology 24: 328‐332. . [BIOLOGY section] Dillon H.F., Somanchi A., Rao K., Jones. P.J.H. (2007) Nutraceutical compositions form microalgae and related methods of production and administration. WO2007/136428 A2. Dvir I., Stark A.H., Chayoth R, Madar Z., Malis Arad S. (2009) Hypocholesterolemic effects of nutraceuticals produced from the red microalga Porphyridium sp in rats. Nutrients 1: 156‐167. Dvir I., Chayoth R., Sod‐Moriah U., Shany S., Nyska A., Stark A.H., Madar Z., Malis Arad S. (2000) Soluble polysaccharide and biomass of red microalga Porphyridium sp. alter intestinal morphology and reduce serum cholesterol in rats. British Journal of Nutrition 84: 469‐476. Fabregas J., Garcia D., Morales E., Domínguez A., Otero A. (1998) Renewal rate of semicontinuous cultures of the microalga Porphyridium cruen turn modifies phycoerythrin, exopolysaccharide and fatty acid productivity. Journal of Fermentation and Bioengineering 86: 477‐481. Griffiths M.J., Harrison S.T.L. (2009) Lipid productivity as a key characteristic for choosing algal species for biodiesel production. Journal of Applied Phycology 21: 493‐507. Guil‐Guerrero J.L., Belarbi E.H., Rebolloso‐Fuentes M.M. (2001) Eicosapentaenoic and arachidonic acids purification from the red microalga Porphyridium cruentum. Bioseparation 9: 299–306. Huheihel M., Ishanu V., Tal J., Malis Arad S. (2002) Activity of Porphyridium sp. polysaccharide against herpes simplex viruses in vitro and in vivo. Journal of Biochemical and Biophysical Methods 50: 189‐200. Rebolloso Fuentes M.M., García Sánchez J.L., Fernández Sevilla J.M., Acién Fernández F.G., Sánchez Pérez J.A., Molina Grima E. (1999) Outdoor continuous culture of Porphyridium cruentum in a tubular photobioreactor: quantitative analysis of the daily cyclic variation of culture parameters. Journal of Biotechnology 70: 271‐288. Rodolfi L., Chini Zittelli G., Bassi N., Padovani G., Biondi N., Bonini G., Tredici M.R. (2009) Microalgae for oil: strain selection, induction of lipid synthesis, and outdoor mass cultivation in a low‐cost photobioreactor. Biotechnology and Bioengineering 102: 100‐112. Sheath R.G. (2003) Red algae. In: Wehr J.D., Sheath R.G. (eds.) Freshwater Algae of North America: Ecology and Classification. Elsevier Science, pp. 197‐224. [BIOLOGY section]
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8.3 Bacillariophyta 8.3.1
Benthic diatoms (Amphora; Amphiprora; Cylindrotheca; Navicula; Nitzschia)
TAXONOMY Amphora sp.
Figure 33 ‐ Amphora coffeaeformis.
Figure 34 ‐ Light microscopic picture of Amphora sp.
Figure 35 ‐ Light microscopic picture of Amphora coffeaeformis.
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SYMBOLS: D, PIV Phylum Class Order Family Genus Species
Bacillariophyta Bacillariophyceae Thalassiophysales Thalassiophysales Amphora
Related species Amphora is a very large and heterogeneous genus. There are 1032 species names in the algae database at present, of which 170 have been flagged as currently accepted taxonomically. A. coffeaeformis, A. coffeaeformis punctata, A. coffeaeformis linea, A. coffeaeformis tenuis, A. coffeaeformis taylori, A. delicatissima, A. delicatissima capitata.
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Amphiprora hyalina
Figure 36 ‐ Light microscopic picture of Amphiprora sp.
Figure 37‐ Light microscopic picture of Amphiprora sp.
Picture from www.plingfactory.de/
Picture from www.mikroskopie‐ph.de/
SYMBOLS: D, PIV Phylum Class Order Family Genus Species
Bacillariophyta Bacillariophyceae Naviculales Amphipleuraceae Amphiprora Amphiprora hyalina
Related species There are 230 species names in the algae database at present, of which 29 have been flagged as currently accepted taxonomically. Synonym: Amphiprora paludosa var. hyalina (Eulenstein ex Van Heurck) Cleve 1894. Cylindrotheca sp.
Figure 38 ‐ Light microscopic picture of Cylindrotheca sp.
Figure 39 ‐ Scanning electron microscopic picture of Cylindrotheca sp.
Picture courtesy of PAE (UGent)
Picture from ocean.inha.ac.kr/l3.htm
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Phylum Class Order Family Genus Species
Bacillariophyta Bacillariophyceae Bacillariales Bacillariaceae Cylindrotheca
Related species There are 6 species names in the algae database at present, of which 2 have been flagged as currently accepted taxonomically. Potentially important for biofuel: • Cylindrotheca fusiformis Reimann & Lewin 1964 • Cylindrotheca closterium (Brébisson ex Kützing) Grunow in van Heurck 1882 Navicula sp.
Figure 40 ‐ Light microscopic picture of Navicula gregaria
Figure 41 ‐ Light microscopic picture of Navicula lanceolata
Picture courtesy of PAE (UGgent)
Picture courtesy of Petr Znachor
SYMBOLS: D, PIV Phylum Class Order Family Genus Species
Bacillariophyta Bacillariophyceae Naviculales Naviculaceae Navicula
Related species Navicula is a very large and heterogeneous genus. There are 6714 species names in the algae database at present, of which 890 have been flagged as currently accepted taxonomically. Potentially important for biofuel Navicula acceptata, Navicula saprophila, Navicula pelliculosa.
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Nitzschia dissipata
Figure 42 ‐ Light microscopic picture of Nitzschia dissipata Picture from http://craticula.ncl.ac.uk/
Figure 43 ‐ Raster electron microscopic picture of Nitzschia sp. Picture from http://ina.beyer‐privat.net/
SYMBOLS: D, PIV Phylum Class Order Family Genus Species
Bacillariophyta Bacillariophyceae Bacillariales Bacillariaceae Nitzschia Nitzschia dissipata
Related species There are 1024 species names in the algae database at present, of which 321 have been flagged as currently accepted taxonomically. Potentially important for biofuel: N. communis, N. frustulum, N. palea.
BIOLOGY Structural and morphological features Benthic diatoms are all raphid, pennate diatoms. Amphora sp. The benthic Amphora species appear epiphytic, epilithic or epipelic (in or attached to sediments). The fouling A. coffeaeformis is a common marine species. Cells of Amphora are solitary, sometimes sessile but usually motile, almost always lying in girdle view and then appearing elliptical or lanceolate, with truncate ends (Round et al., 1990). Amphora species have typical asymmetrical valve morphology. Its dorsiventral frustule resembles ‘‘a third of an orange’’ (Hendey, 1964) with both raphe systems on the same (ventral) side of the cell. Cells usually having 1 or 2, sometimes many, plastids which are extremely diverse in position, shape and structure (Round et al., 1990). Cell length and width varies with species, roughly ranging from 14 – 55 µm and 2.5 – 9 µm respectively (Sala et al., 1998). AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Amphiprora hyalina Amphiprora (currently placed in the genus Entomoneis) is a benthic genus, epipelic of brackish marine sediments; occasionally in freshwater. A. hyalina is a marine species. Amphiprora/Entomoneis cells are solitary and twisted about the apical axis (twisted frustules) usually lying in girdle view and then appearing bilobate. This torsion of the cell means that valves or whole frustules can present a great variety of aspects depending on exactly how they lie relative to the observer. Cells contain one plate‐like plastid or two plastids, one on each side of the median transapical plane. A variety of pore and raphe structures is found in Entomoneis. The raphe is median, sigmoid, on a raised keel forming a wing (Round et al., 1990). Cylindrotheca sp. Cyclindrotheca is benthic and widely distributed in the epipelon (living on (or in) fine sediments) of marine habitats; rarely occurring in freshwater. The genus is very abundant in coastal waters worldwide (Round et al., 1990). Cylindrotheca cells are solitary, long and narrow, needle‐like and only weakly silicified. The frustules are twisted about the apical axis, consequently the cells rotate as they move through the sediment. Cells contain two or many chloroplasts that are plate‐like or discoid (Round et al., 1990). Navicula sp. All Navicula are benthic diatoms. Cells of Navicula sensu lato diatoms have naviculoid (boat‐shaped) cells that may exist singly or in ribbons. Navicula is Latin for "small ship". The valves are symmetrical both apically and transapically, and may have rounded, acute, or capitate ends. The central area is often distinctly expanded. They contain two girdle‐appressed plastids, one on either side of the apical plane. The raphid system is well developed with a raphe on each valve which makes cells highly motile. Nitzschia dissipata Nitzschia comprises both planktonic and benthic species. N. dissipata is a benthic species and was recorded in freshwater (e.g. Aboal et al., 2003; Day et al., 1995; Krammer and Lange‐Bertalot, 1988; Roberts et al.; 2004) as well as in coastal waters (Tadros and Johansen, 1988). Nitzschia is one of the most difficult genera for species identification and many features are only seen by electron microscopy (e.g. Mann, 1986; Trobajo et al., 2004, 2006). Nitzschia cells are usually linear to lanceolate and may be solitary or colony forming. Most species have two chloroplasts, one in each end of the cell. Each valve possesses a raphe that is more or less eccentric and supported by fibulae (=bridges of silica between portions of the valve on either side of the raphe, giving a ladder‐like appearance). The two raphes of a frustule are positioned diagonally opposite (nitzschioid). Valve striae (= lines with small holes) are usually uniseriate. Biofilm formation Benthic diatoms are the most common benthic microalgae, which are abundant in many soft‐sediment aquatic habitats (estuaries, shallow subtidal seas, coral reef flats, lakes, and rivers) and can contribute up to 50% of the total autotrophic production in some ecosystems. They form biofilms, a matrix of cells, sediments and extracellular polymeric substances (EPS) (Underwood and Paterson, 2000). It is known that diatom mucilages are rich in polysaccharides and proteoglycans and are secreted through the channels and pores in the diatom frustule. In some benthic species, these compounds produce structures (tubes, pads and stalks) that are used for attachment to surfaces and often contribute to biofouling problems. Epipelic diatoms do not produce permanent structures but secrete large quantities of extracellular mucilages that are involved in motility (Underwood and Paterson, 2000). Motility is an essential adaptation for photosynthetic organisms in these environments, allowing cells to migrate into the illuminated (photic) zone of sediment near the surface after periods of sediment mixing or deposition. In diatoms, this motility AquaFUELs‐ Taxonomy, Biology and Biotechnology
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is generated by the production of extracellular polymeric substances (EPS), primarily polysaccharides (Underwood et al., 2004). The mechanism of movement in diatoms is unique for microbial cells and relies on the extrusion of mucilage through a slit in the surface of the silica frustule (cell wall). This slit, known as the raphe, may be present on only a single valve of the frustule (monoraphid) or on both valves (biraphid) (Underwoos and Paterson, 2000).The production of EPS in the aquatic environment is ecologically significant because EPS and other carbohydrate‐rich exudates can be used by bacteria, meiofauna, and macrofauna as a carbon source and EPS can increase the stability of sediments (Underwood et al., 2004). The production of extracellular carbohydrates shows some dependency on environmental conditions, for example, irradiance and nutrient conditions. The production of some extracellular carbohydrates ceases in darkness, but studies on axenic cultures of benthic diatoms and natural sediment assemblages have shown production of EPS in dark as well as under illuminated conditions. Continued EPS production in darkness indicates that EPS production is not directly coupled to the photosynthetic production of carbohydrates (Underwood et al., 2004). Nutrient limitation can also increase extracellular carbohydrate production in benthic diatoms: this production of extracellular carbohydrates is assumed to be due to unbalanced metabolism, with cells releasing fixed carbon in excess of their energetic requirements, because of growth being prevented by nutrient (N, P) limitation (Underwood et al., 2004). Diatom EPS consists of a relatively undefined complex mixture of proteins, proteoglycans and carbohydrates (Underwood and Paterson, 2000).There is evidence that benthic diatoms produce a number of different types of EPS, which vary in structure and sugar composition, and that the production of these EPS depends on environmental conditions and the nutrient status of the cells (Underwood et al., 2004). The biosynthetic pathway for these carbohydrates and the mechanisms causing changes in EPS composition are yet to be elucidated. Glucans either may be the precursor of EPS or may act as the photoassimilate carbon store, providing energy for EPS synthesis during periods of darkness.This latter hypothesis is supported by the significant correlation found between glucan catabolism and EPS production in the dark in a number of benthic species. This provides indirect evidence that glucans are involved in the production of EPS (Underwood et al., 2004). Microphytobenthic biofilms can have high rates of photosynthesis and a significant proportion of their photo‐assimilated carbon is released into the environment as extracellular carbohydrates (Underwood and Paterson, 2000). Biochemical composition under optimal and stressed conditions Amphora sp.: The biochemical composition vary among Amphora species as indicated by the diverse values reported by different studies (de la Peña, 2007; Gordon et al., 2006; Khatoon et al., 2009; Sheehan et al., 1998). In addition, different culture conditions result in significant variations in growth and the biochemical composition of the cells of the same strain as shown by de la Peña (2007). He showed that the proximate chemical composition (protein, carbohydrates, fatty acid content, chlorophyll a) of Amphora sp. is highly dependent on light intensity, the culture location (indoor‐outdoor) and the type of enrichment used (de la Peña, 2007). A higher protein and carbohydrate content of Amphora sp. was noted in cultures located inside the laboratory compared to cultures grown outside (probably due to the more regulated cultural conditions inside like constant irradiance and temperature). Lipid content ranged from 26 – 81% depending on culture site and nutrients used (de la Peña, 2007). Amphora is rich in total lipids and fatty acids with a high amount of polyunsaturated fatty acids (PUFAs) especially EPA and a high amount of essential amino acids (Gordon et al., 2006; Khatoon et al., 2009). Griffiths and Harrison (2009) calculated the average total lipid content for Amphora sp. from available literature data. • 51% cdw (cell dry weight) under nutrient replete laboratory conditions • 40% cdw in outdoor ponds
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Amphiprora hyalina: A. paludosa var. hyalina contains a high amount of EPA (28%) (Correa‐Reyes et al., 2009). Proximate composition (dry weight percentage) in A. paludosa var. hyalina (Correa‐Reyes et al., 2009): • 11.33% ± 0.36 protein • 20.96% ± 0.94 nitrogen free extracts • 8.10% ± 0.49 lipids • 59.61% ± 1.01 ash The average total lipid content for Amphiprora hyalina calculated from available literature data by Griffiths and Harrison (2009) is: • 22% cdw under nutrient replete laboratory conditions • 28% cdw under N deficient laboratory conditions • 37% cdw under Si deficient conditions Cylindrotheca sp.: The average total lipid content for Cylindrotheca sp. calculated from literature by Griffiths and Harrison (2009) is: • 27% cdw under nutrient replete conditions • 27% cdw under N deficient conditions Navicula sp.: The average total lipid content for N. acceptata calculated from literature by Griffiths and Harrison (2009) is: • 33% cdw under nutrient replete conditions • 35% cdw under N deficient conditions • 46% cdw under Si deficient conditions N. saprophila contains large quantities of EPA and is considered a potential source of this important fatty acid (Kitano et al., 1997). It was shown that EPA production was enhanced under mixotrophic conditions in CO2 enriched (about 2%) atmosphere in the presence of acetate as compared with photoautotrophic conditions (Kitano et al., 1997; 1998). The biomass of a freshwater N. saprophila strain has the following composition (Pilny, 2009): • • • •
Protein Lipid Crude fiber Ash
grown in WC medium 46.03 % 28.57 % 34°C. Water hyacinth is reported to tolerate grazing. Native to Brazil, now growing in most tropical and subtropical countries. More than 50 countries in which water hyacinth is weed have been listed. It grows in ponds, ditches, canals, calm waters of rivers at an elevation of 0‐300 m and a depth range of 2 m. Engineers have estimated that the Panama Canal would be impassable within three years without continuous aquatic weed control measures. Fresh plant contains 95.5% moisture. On a dry matter basis, it is 75.8% organic matter, 1.5% N, and 24.2% ash. The ash contains 28.7% K2O, 1.8% Na2O, 12.8% CaO, 21.0% Cl, and 7.0% P2O5. Protein is bout 12‐ 18% of dry matter. Protein contains, per 100 g, 0.72 g methionine, 4.72 g phenylalanine, 4.32 g threonine, 5.34 g lysine, 4.32 g isoleucine, 0.27 g valine, and 7.2 g leucine. 23‐28% of dry weight is cellulose. On a dry matter basis, standing crop is 12.8 t ha‐1, maximum productivity 14.6 g m‐2 day‐1, maximum yield 54.7 g m‐2 day‐1.
BIOTECHNOLOGY Water hyacinth would be ideally suited for nutrient removal systems. Aquatic plants, when growing in water containing ample quantities of N, P and K, will exploit the situation by “luxury consumption” of these elements, far in excess of what they need for healthy growth. As it floats on the surface and is not rooted, harvesting is facilitated. By continuous harvesting the population could be kept in a rapidly expanding phase during which uptake rates of nutrients are at their highest. Waters beneath dense stands are anaerobic so additional N would be lost by denitrification. There would be considerable microbial activity beneath the hyacinths and nutrients would be absorbed by these organisms. However, considerable organic matter would reach the water by the loss of root fragments that probably have a fairly high biological oxygen demand (BOD) and it might prove necessary to use conventional sewage holding ponds to reduce the BOD prior to final release. Water hyacinth roots naturally absorb pollutants, including such toxic chemicals as lead, mercury, and strontium 90, as well as some organic pollutant, in concentrations 10,000 times that in the surrounding water. Water hyacinths have been used as human food in the Philippines in war time. The soft white bud of the plant was eaten either raw, or as a salad, or as an ingredient in vegetable dishes. It was called ‘repollo’ (cauliflower). Under conditions of relative food abundance it is considered unlikely that it would still be used as food. Dried and cleansed plants, can be used as fertilizer, poultry feed, additives to cattle‐feed, and plant mulch. Water hyacinth protein shows deficient levels in only two of the essential amino acids, valine and methionine, as compared to the FAO reference pattern. A diet containing an adequate protein level will be balanced in lysine if it contains 4.2% of that amino acid in its protein content. Corn is deficient in lysine, containing only 0.8%. However water hyacinth contains 5.3% lysine, while milk contains 7.8%. It is evident that water hyacinth could serve to improve the lysine content of a corn diet. Leaf protein which contains 6.3% lysine has been reported to be an effective supplement for barley in pig rations. As a feed for ruminants, water hyacinth should be administered as a combination of the lamina , where the protein and fibre mainly are, and the petiole where the carbohydrate are mainly located. Animals eating fresh water hyacinth as sole ingredient of the diet could not ingest sufficient dry matter, quite apart from the imbalance of the nutrients. On a dry matter basis water hyacinth is better than straw but a little low in protein to compare with hay. The physical structure of the plant is not suitable for hay or silage making and the product would not have much nutritional value. The nutritional value could be increased by mixing with molasses. The hyacinth is rich in minerals, but it would be simple to incorporate any necessary food AquaFUELs‐ Taxonomy, Biology and Biotechnology
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additives in the processing. Pig farming using water hyacinth grown in ponds fertilized with human and animal waste is well established in Malaysia. The hyacinth can be fed fresh after removal of the roots. Some people boil the plant. Pigs readily eat the hyacinth and thrive on it. Some farmers have machines for slicing the plants into small pieces. The high vitamin content (A, B and C) and easy availability of the plant are important advantages. But the high moisture content and the possibility of the material being contaminated with pathogens seem to deter the extensive practice of fresh plant feeding. Consequently most farmers (70–80%) prefer to cook the sliced hyacinth with other feed ingredients for 5 to 6 hours and feed the boiled material. Silage is also fed to the pigs. Good compost can be made from hyacinth. Attempts have been made to utilize the plant as a raw material for paper, plastics and other commercial products, but so far no industry based on water hyacinth appears to have been established. The fibrous stem, constituting about 40% of the whole plant, is suitable for paper making. The addition of jute or cotton fibres to the extent of 8–10% on the weight of the pulp is considered necessary as the paper prepared from the stems alone is translucent. A plastic material suitable for the production of moulded articles and boards has been prepared from water hyacinth. In Asia, the plants are collected at the beginning of the cold weather, left to dry and the dry material used along with jute sticks as fuel. The ashes are subsequently used as manure. The possibility of using the dried weed for the production of power gas and power alcohol has been considered. Three methods have been suggested, viz. saccharification by acid digestion and subsequent fermentation, gasification by air and steam with recovery of ammonia, and bacterial fermentation and utilization of the evolved gas for power production. Potassium chloride (0.1 ton of KCl per ton of dry hyacinth) is recovered in all the processes. Starting from 1 ton of dried water hyacinth, 59 L of ethyl alcohol and 0.2 tons of residual fibre (8.1 MJ) are obtained by the first process. Gasification by air and steam gives, per ton of dried material, 37‐ 53 kg of ammonium sulphate and 1133 m3 of gas (0.16 MJ) containing hydrogen, 16.6%; methane, 4.8%; carbon monoxide, 21.7%; carbon dioxide, 4.1%; and nitrogen, 52.8%. Bacterial fermentation gives per ton of material 750 m3 of gas (0.63 MJ) containing: methane, 51.6%; hydrogen, 25.4%; carbon dioxide, 22.1%; and oxygen, 1.2%. The commercial possibilities of the processes have not been proved. References Anonymous (1976) Water hyacinths soak up pollution. BioScience 26: 234. [BIOTECHNOLOGY section] Bailey L.H. (1949) Manual of Cultivated Plants. Macmillan, New York. Duke J. Handbook of energy crops. http://www.hort.purdue.edu/newcrop/duke_energy/dukeindex.html [BIOTECHNOLOGY section] Gohl B. (1981) Tropical feeds. Feed information summaries and nutritive values. FAO Animal Production and Health Series 12. FAO, Rome. [BIOLOGY section] Little E.C.S. (1979) Handbook of utilization of aquatic plants. FAO Fisheries Technical Paper No. 187, Rome. [BIOTECHNOLOGY section] Hammer R.L. (1996) Eichhornia crassipes. In Randall J.M., Marinelli J. (eds.) Invasive Plants: Weeds of the Global Garden. Brooklyn Botanic Garden Inc., New York, p. 99. [BIOLOGY section] Holm L.G., Plunknett D.L., Pancho J.V., Herberger J.P. (1977) The world's worst weeds. University Press of Hawaii, Honolulu. [BIOLOGY section] Holm L.G., Pancho J.V., Herberger J.P., Plucknett, D.L. (1979) A geographical atlas of world weeds. John Wiley & Sons, New York. [BIOLOGY section] Integrated Taxonomic Information System (ITIS) catalogue of life 2010 http://www.catalogueoflife.org/annual‐ checklist/2010/search/all/key/eichhornia/match/1 (accessed on the 10th of June 2010) [BIOLOGY section] Matai S., Bagchi D.K. (1980) Water hyacinth: a plant with prolific bioproductivity and photosynthesis. In: Gnanam A., Krishnaswamy S., Kahn J.S. (eds.) Proceedings of the International Symposium on Biological Applications of Solar Energy. MacMillan Co. of India, Madras, pp. 144‐148. [BIOLOGY section] Penfound W.T., Earle T.T. (1948) The biology of the water hyacinth. Ecological Monographs 18: 449‐472. [BIOLOGY section] Reed C.F. (1970) Selected weeds of the United States. Agriculture Handbook 366. USDA, Washington, DC. [BIOLOGY section] USDA, NRCS( 2001) The PLANTS Database, Version 3.1. (http://plants.usda.gov). National Plant Data Center, Baton Rouge, LA 70874‐4490 USA.
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10.3 Elodea canadensis
Figure 93 ‐ Elodea canadensis Michaux (1803). Illustration by C. A. M. Lindman
SYMBOLS: B
TAXONOMY
Phylum Class Order Family Genus Species
Magnoliophyta Liliopsida Hydrocharitales Hydrocharitaceae Elodea Elodea canadensis
Related species E. bifoliata, E. callitrichoides, E. granatensis, E. nuttallii, E. potamogeton, E. spinosa.
BIOLOGY Elodea canadensis is a submerged aquatic perennial freshwater herb, usually firmly rooted to the bottom mud and producing a thick green mat below the water surface. Short thread like stolons giving rise to slender vertical stems to 3 m long. Leaves are dark green oblong‐linear, formed at intervals of 3‐25 mm along the stem, in groups of three, each 6‐12 mm long and about 1‐5 mm wide, usually with forward pointing teeth on the margins. Roots are filamentous, rising from the nodes along the stolons. Flowers are white or pale purple with three sepals and petals. They are solitary, forming in the axils of the leaves and growing towards the surface on threadlike stalks about 30 cm long.. Fruits are capsules less than 1 cm in length. It is dispersed by seeds and fragments via water currents. It is a dioecious plant flowering from June to August. Pollination occurs near the water surface and pollen is distributed by wind and water currents. Vegetative reproduction by fragments is very common. Mass development has been reported multiple
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times in the last century. It is preyed upon by a high number of freshwater organisms including fish and birds. It is diffuse in surface standing waters, and surface running waters, like shallow lakes, ponds, pools, ditches and streams with slow moving water. It grows up to 3 m water depth and, in exceptional cases, up to 16 m in depth. It tolerates pH values from 6.0 to 7.5 and temperatures from 1 to 25 °C. It originates from North American inland waters. The first European record was reported from Ireland in 1836. It subsequently became widespread in north and central European countries. After a rapid colonization of northern Europe the populations declined due to the introduction of Elodea nuttallii. Today the population is stable. As it can be very dominant, it competes for nutrients and space with other plants. It can bioaccumulate nutrients and modify the habitat by reducing water movement. The species is known to outcompete other plants. During dense blooms, impairs boating, fishing, swimming, and water skiing. Clogging of water intake pipes of power plants and other industries were reported. Elodea is anchored to the bottom deposits of a lake or river by adventitious roots, but nutrients are probably absorbed mainly by leaves and stems in contact with the free water. Although the concentration of dissolved salts in the surrounding water will largely control the density and composition of the plant, the evidence indicated a difference in composition between species, even in the same lake, as well as seasonal changes. It follows that if this water weed is to be exploited commercially, preliminary plant analyses are needed to determine the best species and season for harvesting. Crude protein is about 12%, ash 28% and fiber 16%.
BIOTECHNOLOGY To obtain 1 ton of dry Elodea canadensis 14 tons of wet material would have to be processed. E. canadensis is an excellent food for cattle and pigs, when fed to sheep it was found to be unpalatable but it was accepted when mixed with pasture (1:5 dry matter basis). It appears to contain all the vitamins in at least as high an amount as alfalfa. Of more interest is the fact that the biological value of the protein in Elodea is about 70% that of alfalfa and the digestibility is apparently better. Cystine seems to be the limiting amino acid in Elodea as well as in alfalfa. E. canadensis can be used as a non‐expensive supplemental food in order to increase growth and survival in summerling noble crayfish, A. astacus, that has the potential to consume this macrophyte in nature. It shows the typical productivity of submerged macrophytes in temperate zones as 2–7 tons dry weight ha‐1 year‐1 and 17–59 tons fresh weight ha‐1 year‐1. References Bowmer H., Kathleen S.W., Jacobs L., Sainty G.R. (1995) Identification, biology and management of Elodea canadensis, Hydrocharitaceae. Journal of Aquatic Plant Management 33: 13‐19. [BIOLOGY section] Champion P.D., Hofstra D.E., Clayton J.S. (2007) Border control for potential aquatic weeds. Stage 3. Weed risk management. Science & Technical Publishing New Zealand Department of Conservation. [BIOLOGY section] D’Agaro E., Renai B., Gherardi F.(2004) Evaluation of the American waterweed (Elodea canadensis Michx.) as supplemental food for the noble crayfish, Astacus astacus. Bulletin Français de la Pêche et de la Pisciculture 372‐373: 439‐445. [BIOTECHNOLOGY section] Department of Agriculture and Food, Australia: http://agspsrv95.agric.wa.gov.au/dps/version02/01_plantview.asp? [BIOLOGY section] Gollasch S. (2006) Elodea canadensis. DAISIE‐Delivering Alien Invasive Species Inventories for Europe. http://www.europe‐ th aliens.org/pdf/Elodea_canadensis.pdf (accessed on the 10 of June 2010) [BIOLOGY section] Integrated Taxonomic Information System (ITIS) catalogue of life 2010 http://www.catalogueoflife.org/annual‐ checklist/2010/search/all/key/elodea/match/1 (accessed on the 10th of June 2010) [BIOLOGY section] th Kews World Checklist of Selected Plant Families http://apps.kew.org/ (accessed on the 10 of June 2010) [BIOLOGY section] Little E.C.S. (1979) Handbook of utilization of aquatic plants. FAO Fisheries Technical Paper No. 187, Rome. [BIOTECHNOLOGY section]
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10.4 Lagarosiphon major
Figure 93 ‐ Lagarosiphon major (Ridley) Moss © Control of aquatic invasive species in Ireland (CAISIE Life+ project)
SYMBOLS: B
TAXONOMY
Phylum Class Order Family Genus Species
Magnoliophyta Liliopsida Alismatales Hydrocharitaceae Lagarosiphon Lagarosiphon major
Related species L. cordofanus, L. hydrilloides, L. ilicifolius, L. madagascariensis, L. muscoides, L. rubellus, L. steudneri, L. verticillifolius.
BIOLOGY L. major is a rhizomatous, perennial, submerged aquatic plant. It reaches its maximum growth in clear water up to a depth of 6.5 m, but may only grow to 1 m in murky water. It has numerous threadlike roots, which are adventitious and, along with rhizomes, anchor it to the bottom. Stems, which can reach the surface, are brittle and sparsely branched, 3‐5 mm in diameter and curved towards the base (J‐shaped). The leaves are 5‐20 mm long and 2‐3 mm wide, and occur in alternate spirals along the stem. They generally have tapered tips curving downwards towards the stem, except in low alkalinity water where they are straight. The three‐petalled female flowers are very small, clear‐white on the surface, and grow on very thin white to almost translucent filament‐like stalks. Neither the male flower, which floats freely to the AquaFUELs‐ Taxonomy, Biology and Biotechnology
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surface, nor fruit or seeds have been recorded outside of its native range. Since the species is dioecious (sexes on different plants) both must be present for sexual reproduction. Only female plants are known outside of the native range of this species. All reproduction in introduced regions is therefore asexual primarily by fragmentation or local growth by rhizomatous spread. L. major grows best in clear, still or slow‐moving fresh water with silty or sandy bottoms. It prefers the cooler waters of the temperate zone, with optimum temperatures of 20‐23 °C and a maximum temperature of around 25 °C. It can live in high and low nutrient levels and grows best under conditions of high light intensity. It also tolerates relatively high pH (i.e. alkaline conditions). Growth of L. major is greatest in sheltered areas protected from wind, waves and currents. It is native of southern Africa, is found in high mountain streams and ponds. It has spread throughout the world as an aquarium plant and is also known as an 'oxygen plant'. However, dense infestations can actually consume more oxygen than they produce, and reduce water quality and available oxygen.
BIOTECHNOLOGY L. major and other aquatic species grown in small outdoor tanks can be used successfully to assess the effects of crop‐protection products on non‐target aquatic flora. The possible utilization by harvesting for stock feed in New Zealand lakes was evaluated; however, the use of the plants as fodder was thought to be unsuitable because of the content of arsenic accumulated by the plants from the thermal waters that enter the lakes. References Airo S., Sconfietti R. (1995) In situ experiments on productivity of aquatic macrophytes in a pond. Rivista‐di‐Idrobiologia 34: 147‐ 156. CAISIE life+project (2010) control of aquatic invasive species in Ireland. www.caisie.ie. Accessed on the 06th of October 2010. Champion P.D., Tanner C.C. (2000) Seasonality of macrophytes and interaction with flow in a New Zealand lowland stream. Hydrobiologia 441: 1‐12. Coffey B.T., Clayton J.S. (1987) Submerged macrophytes of Lake Pupuke Takapuna New Zealand. New‐Zealand Journal of Marine and Freshwater Research 21: 193‐198. CONABIO (2008) Sistema de información sobre especies invasoras en México. Especies invasoras ‐ Plantas. Comisión Nacional para el Conocimiento y Uso de la Biodiversidad. Fecha de acceso. www.conabio.gob.mx/invasoras/index.php/Especies_invasoras_‐_Plantas Conservatoire Botanique National De Mascarin (BOULLET V. coord.) (2007) Lagarosiphon major. Index de la flore vasculaire de la Réunion (Trachéophytes) : statuts, menaces et protections. ‐ Version 2007.1 (mise à jour 12 juin 2007). Cook C.D.K. (2004) Aquatic and Wetland Plants of Southern Africa. Backhuys Publishers, The Netherlands. de Carvalho R.F., Bromilow R.H., Greenwood R. (2007) Uptake of pesticides from water by curly waterweed Lagarosiphon major and lesser duckweed Lemna minor. Pest Management Science 63:789–797. [BIOTECHNOLOGY section] Egloff F. (1975) New and noteworthy species of Swiss flora. Bulletin de la Societe Botanique Suisse 84: 333‐342. Global Invasive Species Database: http://www.issg.org/database/species/ecology.asp?si=403&fr=1&sts=sss [BIOLOGY section] and [BIOTECHNOLOGY section] ITIS (Integrated Taxonomic Information System) (2005) Online Database Lagarosiphon major. http://www.itis.gov/servlet/SingleRpt/SingleRpt?search_topic=TSN&search_value=565981 James C.S., Eaton J.W., Hardwick K. (1999) Competition between three submerged macrophytes, Elodea canadensis Michx, Elodea nuttallii (Planch.) St John and Lagarosiphon major (Ridl.) Moss. Hydrobiologia 415: 35‐40. Little E.C.S. (1979) Handbook of utilization of aquatic plants. FAO Fisheries Technical Paper No. 187, Rome. [BIOTECHNOLOGY section] Rattray M.R. (1995) The relationship between P, Fe and Mn uptakes by submersed rooted angiosperms. Hydrobiologia 308: 117‐ 120. Rattray M.R., Howard‐Williams C., Brown J.M. (1994) Rates of early growth of propagules of Lagarosiphon major and Myriophyllum triphyllum in lakes of differing trophic status. New Zealand Journal of Marine and Freshwater Research 28: 235‐241. Riis T., Biggs B.J., Flanagan M. (2003) Seasonal changes in macrophyte biomass in South Island lowland streams, New Zealand. New Zealand Journal of Marine and Freshwater Research 37: 381‐388. Roy B., Popay I., Champion P. James T., Rahman A. (2004) An Illustrated Guide to Common Weeds of New Zealand 2nd Edition. Lagarosiphon major oxygen weed. New Zealand Plant Protection Society. State of Queensland (2004) Lagarosiphon major description. The State of Queensland (Department of Natural Resources and Mines). Strickland R., Harding J., Shearer L. (2000) The Biology of Lake Dunstan. Cawthron Report No. 563; Contact Energy Limited.
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Symoens L., Triest S. (1983) Monograph of the African genus Lagarosiphon. Bullettin du Jardin Botanique National de la Belgique 53: 441‐488. University of Florida (2001) Lagarosiphon major (Ridley) Moss. Non‐Native Invasive Aquatic Plants in the United States, Center for Aquatic and Invasive Plants, University of Florida and Sea Grant. Wells R.D., De‐Winton M.D., Clayton J.S. (1997) Successive macrophyte invasions within the submerged flora of Lake Tarawera, central North Island, New Zealand. New Zealand Journal of Marine and Freshwater Research 31: 449‐459.
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10.5 Lemna minor
Figure 95 ‐ Lemna minor Linnaeus Robert H. Mohlenbrock @ USDA‐NRCS PLANTS Database / USDA NRCS. 1995. Northeast wetland flora: Field office guide to plant species. Northeast National Technical Center, Chester
SYMBOLS: B, E, PIV
TAXONOMY
Phylum Class Order Family Genus Species
Magnoliophyta Liliopsida Arales Lemnaceae Lemna Lemna minor
Related species L. aequinoctialis, L. disperma, L. gibba, L. japonica, L. minuta, L. obscura, L. perpusilla, L. tenera, L. trisulca, L. turioni fera, L.mna valdiviana, L. yungensis.
BIOLOGY General Description of Major divisions and classes: The Lemnaceae is a monocotyledonous family of 4 genera: Spirodela, Lemna, Wolffia and Wolfiella, and 37 species. All Lemnaceae species are small aquatic plants, commonly called duckweeds (Lemna and Spirodela species) and water meals (Wolffia species). The majority of research involving these plants has been done with only a few species; primarily L. gibba and L. minor, Spirodela polyrrhiza and Spirodela punctata, and to a lesser extent, Wolffia globosa. For the majority of species, little is known of their biology and the ability to extrapolate any of the technological applications developed for the better studied species (Stomp, 2005). AquaFUELs‐ Taxonomy, Biology and Biotechnology
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Physiological characteristics
The plants are perennials, growing anywhere there is water and sun. These plants require little mechanical support or vascular tissue (the smallest members of the family completely lack xylem and phloem) so most cells resemble maturing leaves expending very little photosynthetic energy on plant structures. As a result, vegetative propagation of these plants has high biomass yield potential. Structural and morphological features Members of the family Lemnaceae are tiny aquatic monocots that range in size from 1.5 cm long (Spirodela polyrhiza) to less than a millimeter (Wolffia globosa). All members of the Lemnaceae are small, free‐floating, fresh‐water plants whose geographical range spans the entire globe (Landolt, 1986). Members of the Lemnaceae are the most morphologically reduced plants known. Plants of Spirodela and Lemna species consist of a frond, a root or roots (the number of roots are species specific) and, when present, a flower. The morphology of Wolffia species is further reduced, with plants consisting of tiny fronds devoid of roots and producing single flowers. Wolfiella species are more varied in morphology. Biochemical composition Table 21 ‐ Biochemical composition of Duckweed. Composition expressed as % dry weight except for Water and ash. From Landolt and Kandeler (1987a), except for starch (Cheng and Stomp, 2009). Water (%FW) Ash Total Carbohydrate Crude fiber Starch Protein Lipid K Na Ca Mg Cu Si S P
Duckweed (Lemnaceae) 86‐97 12.0‐27.6 14.1‐43.6 5.7‐16.2 6‐75 (some strains in lab condition) 6.8‐ 45.0 1.8 ‐9.2 0.03‐7.0 0.03‐1.3 0.18‐4.5 0.04‐2.8 0.2‐10‐3 ‐ 3.2 0.41‐5.35 0.33‐7.0 0.03‐2.8
The protein content of duckweeds is one of the highest in the plant kingdom, but it is dependent on growth conditions. Typically duckweeds are rich in leucine, threonine, valine, isoleucine and phenylalanine. They tend to be low in cysteine, methionine, and tyrosine. Duckweed growth can be optimized to produce high levels of protein or high levels of starch. The plant's dry weight accumulation varies by species and growth conditions and ranges from 6 to 20% of fresh weight (Landolt and Kandeler, 1987b; Tillberg, 1979). Protein content of a number of duckweed species grown under varying conditions has been reported to range from 15 to 45% dry weight (Chang, 1977; Porath, 1979; Appenroth, 1982). These values place the protein content of dry duckweed biomass between alfalfa meal (20%) and soybean meal (41.7%) (Hillman, 1961). We routinely grow duckweed on dilute swine wastewater and get 30 to 35% protein of dry duckweed. Duckweed starch content is dependent on growth conditions, e.g., pH, phosphate concentration (Tasseron‐De‐Jong, 1971; McLaren and Smith, 1976) and
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developmental states con‐ trolled by the plant hormones, cytokinin (Tasseron‐De‐Jong, 1971; McCombs and Ralph, 1972) and abscissic acid (Landolt and Kandeler, 1987b; McLaren and Smith, 1976). Starch contents ranging from 3 to 75% have been reported (Landolt and Kandeler, 1987b; Reid and Bieleski, 1970). A duckweed starch content of 75% is comparable to corn, whose starch content ranges from 65 to 75% (Lin and Tanaka, 2006). Growth kinetics and productivity Doubling times vary by species and environmental conditions and are as short as 20 to 24 hours and many species have doubling times of 2 to 3 days (Chang et al, 2003; Venkararaman et al, 1970). Intensive laboratory culture of duckweed has achieved high rates of biomass accumulation per unit time at culture densities of 1–2 kg m‐2 (Landolt and Kandeler, 1987b). In wastewater treatment research, a growth rate of 0.2 kg dry weight m‐2 week‐1 has been achieved (Cheng et al., 2002a). To achieve these growth rates, only low concentrations of nutrients are required. Oron and co‐workers (1988) achieved optimal growth rates at 20 ppm nitrogen utilizing municipal waste‐ water. Our research with wastewater indicates that high growth rates can be achieved at nitrogen levels less than 10 ppm (Cheng et al., 2002a, b).
BIOTECHNOLOGY Culture media Stomp (2005) and others have demonstrated duckweed growth on a variety of nutrient solutions listed in Table 22. Although differences in growth rates have been observed, generally, duckweed will grow on almost any dilute, inorganic salt solution that supplies essential macro‐ and micro‐nutrients. The plants tolerate a range of pH, for most species ranging between pH 4.5 and 7.2. A number of organic buffers, e.g. EDTA, citrate, tartaric acid, MES, MOPS, and compounds which stabilize proteins, such as PVP, can be added to the growth medium without significantly affecting growth rates. This is an important consideration if recombinant proteins are secreted and are to be recovered from plant growth medium. If the plants are grown under light levels insufficient to support robust photosynthetic growth (
