Why is it common to grow microalgae in bottles or canisters?

Why is it common to grow microalgae in bottles or canisters?

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I came across many people growing Chlorella or Spirulina types of microalgae, inside bottles or canisters and that that they also bubbled CO2 into these containers artificially.

This question is basically comprised of two questions that will clarify what I am trying to understand:

  • Why do they use bottles or any type of bottle-like canister?
  • Why do they insert CO2 to the bottle artificially in cases the bottle doesn't have a cork ? If the claim is that enough CO2 won't be able to get through the bottle neck than why growing the algae inside a bottleaquarium in the first place, and not in a tub with full exposure to the air?

Really, the credit for this answer goes to another 'Homo sapien' from the Spirulina manual link in his comment.

Why do they use bottles or any type of bottle-like canister?

Likely, because they are clear. From the manual:

Spirulina needs sunlight so it is preferable that the container in which it is grown be transparent.

Plastic bottles are cheap, clear, and easy to come by, so they make the ideal material.

Why do they insert CO2 to the bottle artificially in cases the bottle doesn't have a cork?

I don't think the tubes in your picture are CO2 tubes. Rather, I'd bet they are bubble tubes (like in aquariums). These are necessary to keep the growth medium moving constantly. Again, from the manual:

Spirulina tends to gather at the top of the growing culture, where sunlight exposure is maximal. Due to this, Spirulina that cannot reach the top will not multiply and will ultimately die.

In order to maximize Spirulina exposure to sunlight, the water in which it is grown must be stirred…

Another option is a pump, the simplest kind used for aquariums. (emphasis mine)

Blooming prospects?

The seas swarm with them, an unimaginable multitude. Algae are the most important carbon sink in the biosphere the world’s oceans sequester annually 2 gigatons of carbon via carbon dioxide uptake compared with 1.4 gigatons for the total land biosphere. Yet of the 40 000 or so species of algae that exist, but a tiny number𠅊s few as five or six𠅊re exploited by humans. That minuscule fraction, however, amounts to a consumption of 5਋illion tons per year, mostly marine multicellular green and green-brown algae, or ‘seaweed’. The Japanese eat it every day as ‘nori’, thin sheets of dried seaweed that wrap sushi rolls and accompany the traditional breakfast many westerners probably use it daily on their faces in cosmetics and in common household products (Table ​ (TableI I ).

Instant puddings
Ice cream
Shaving cream
Fertilisers and soil

These multicellular ‘macroalgae’ provide three principal extracts: alginates, carageenins and agars, which are valued for their physical properties, such as thickening, gel-forming, emulsifying or film-forming. But now the time has come for microalgae, unicellular blue-green and red algae that can be grown in vast photosynthetic bioreactors. They are prized for their micronutritional value, containing substances that those who invest in them believe will boost our immune system, fight cancer, protect us from UV radiation and treat joint ailments, to mention but a few applications. Others have explored their use in bioremediation—the removal of heavy metals from polluted soil and water𠅊nd some have even tried to grow microalgae in the CO2-rich flue gas of industrial plants to use the resulting biomass as combustible fuel. But is mass production of microalgae in autotrophic—light-driven𠅋ioreactors economically viable?

Clearly some believe it is because they have built the world’s largest closed photobioreactor, reliant solely on light as the source of energy, on the outskirts of Klötze in northern Germany. With a capacity of 700 cubic metres, and covering an area of more than one hectare, the German green machine is 10 times larger than its nearest competitor in Hawaii. Its product, Chlorella vulgaris, a single-celled blue-green alga, is destined mainly for food and feed additives, and cosmetics.

But a leading authority on microalgae, Michael Melkonian, professor at the Botanical Institute, University of Cologne, thinks there are still some hurdles to overcome. ‘The biggest problem is how to get the source of energy to the cells. To get light to every cell, you need a thin layer of cells higher plants solved that problem millions of years ago with the leaf.’ Melkonian believes that although production of microalgae is certainly a worthy pursuit, the present bioreactors need optimising before they can be economically viable.

This is bad news for Gottfried Mende, director of the Klötze plant, though he remains convinced of the success of his venture, and in particular the value of his product. So full of good things are they, that ‘it is possible to exist purely on Chlorella vulgaris’ according to Mende, 𠆋ut it’s not much fun’ he added humorously. His company, Ökologische Produkte Altmark GmbH, has staked its future on the demand for the green gold, which contains more protein per gram of dry mass than soya, and promises to spawn a range of functional or designer foods. Indeed, last year in Germany alone up to 350 tons dry mass of microalgae were used, mainly in human food additives and cosmetics the Klötze plant produces 150 tons annually in its 500 km of naturally-lit piping, and sells its product for 50 Euros per kg dry mass. However, unlike Soya, it is not for its protein value that Chlorella is eaten by humans and fed to livestock. Though the mechanism is not well understood, a daily dose of 3 g of Chlorella vulgaris can boost the immune system, and make you feel better, says Mende, who takes a dose every day himself.

The boost that Chlorella gives to the immune system is likely to be due to branched chain polysaccharides, which are known to be antigenic the cardiovascular benefits derive mainly from antioxidants and omega-unsaturated fatty acids. These are the very same that are found in some fish—which acquire them by eating algae. Algal antioxidants such as carotenoids and complex polyphenols, soak up toxic radicals, and can protect against cancer, UV damage and atherosclerosis. The public can now buy Mende’s brown-green Chlorella vulgaris tablets in the local pharmacy, and add them to their morning selection of health and vitality-promoting vitamin and mineral supplements.

What is good for man also turns out to be good for beast. Trials have shown that adding 0.2% Chlorella vulgaris to chicken feed increases the animals’ weight at slaughter by 10%. The chickens are generally healthier, and benefit from a 16% reduction in cholesterol. Here the market for microalgae has recently been boosted by the EC’s new regulations limiting the use of feed antibiotics. Moreover, it would appear that there are no risks to humans or the environment from microalgae, since they have inhabited the earth for billions of years, and have been in the human food chain since the dawn of mankind.

Our historical familiarity with algae has made us unaware of their ubiquity thinks Otto Pulz from the Institut für Getreideverarbeitung GmbH near Potsdam, Germany: ‘I think that algae are already more public than the public realises,’ remarked Pulz, who works on photobioreactor development, and use of microalgal extracts in cosmetics. The consumer acceptance of algal products has increased dramatically in the last 5 years according to Pulz, who is a key figure in the development of the plant in Klötze. And because of the enormous biodiversity in the oceans, bioengineers need only look to nature to find new promising compounds or biochemical pathways. ‘Genetic engineering in microalgae is not a real aim for the biotechnologist,’ as Pulz put it. The potential applications of natural microalgal extracts are growing almost daily (Table ​ (TableII II ).

Medicinal and general health:
Stimulation of a vigorous immune system
Treatment of carpal tunnel syndrome
Treatment of joint pain and inflammation
Fluorescent pigments for medical diagnosis
Improved muscle recovery after exercise
Treatment of cold sores
Treatment of allergies
Cancer treatment
Increased general health in humans and farm animals
Cardiovascular disease prevention
Cancer prevention
UV protection

Entrepreneurs across the Atlantic also believe in the economic viability of large-scale microalgae production. If microalgae are being touted as the next functional food additive in Germany, this can be nothing compared with the hype that surrounds their health-promoting qualities in the USA. Idyllically situated, the waterfront complex in Kailua-Kona, Hawaii, is the largest open culture facility in the world. Glistening emerald green in the pacific sun are 36 hectares of interlinked channels containing Spirulina pacifica, a spiral microalga that thrives at pH 10�. The company is Cyanotech, and its CEO, Gerry Cysewski certainly believes he is onto a winner. Spirulina Pacifica ® —now a registered trademark—is presented as 𠆊 nutrient-rich dietary supplement… ֺ] highly absorbable source of natural beta-carotene, mixed carotenoids and other phytonutrients, B vitamins, gamma linolenic acid (GLA), protein and essential amino acids.’ Amazingly this is also true, but it may come as a surprise to western cultures raised on synthetic food that anything so natural could possibly be good for you. Indeed, according to Cyanogen, it ‘has been used as a significant food source for centuries.’

Ironically we have rediscovered via science the accepted wisdom that plants contain a great deal of good things. Nevertheless, brownish tablets will hardly inspire the consumer, so Cyanotech markets it in attractive colourful bottles. Apparently six Spirulina tablets (0.5 g each) provide the essential nutrients of 5𠄷 servings of fruit and vegetables recommended by the US cancer and heart societies. To add weight to its claimed health benefits, Spirulina has been shown recently to increase the tumor killing ability of Natural Killer cells and interferon gamma, according to a clinical trial on volunteers at the Osaka Institute of Public Health in Japan.

And this may not be the only clinical application of microalgae. Another extract may soon be prescribed for carpal tunnel syndrome and joint inflammation. As the Cyanotech CEO explained, natural astaxanthin—trademark BioAstin‘𠅊 pigment and antioxidant produced by the alga Haematococcus, is the most promising treatment for carpal tunnel syndrome short of surgery. In the USA alone the market for carpal tunnel syndrome and joint ailments is US$ 1.5਋illion annually. Produced in a 45਌ubic metre closed photobioreactor at Cyanotech, astaxanthin also claims to be beneficial against muscle soreness and reduced immunity resulting from free radical release during strenuous exercise. But the success of natural remedies also depends on the science behind the claims. As Cysewski pointed out, ‘Many products in the US health industry have a lot of questionable science behind them’ this is why Cyanotech is keen to ‘use sound science to support health claims’. The plant also employs environmental practise, recycles all its water and avoids the use of pesticides or herbicides.

After all, microalgae, and plants in general, are well known to be among the world’s greatest environmental cleaning devices. In fact, the prospect that microalgae could be used in environmental bioremediation has been considered for decades. Pilot studies have been conducted in Germany and the USA on the feasibility of using microalgae to remove carbon dioxide from the flue gases of industrial plants, and to use the resulting biomass as combustible fuel. They might also find applications in the removal of heavy metal contamination from water and soil dentists in Germany already give patients Chlorella tablets to absorb the mercury released during the replacement of mercury amalgam fillings.

But tempting though it is to believe that microalgae could be the solution to global CO2 overproduction, Michael Melkonian can speak of frustrating experiences to the contrary. As a consultant to several industrial projects, he should not be ignored. Working on a 1970s project to reduce CO2 emissions from a Westfalian brown coal plant have taught him that algae are not an economically feasible solution. Though the algae will grow like wildfire in 10% CO2 they must be produced at a cost of no more than 40 Euros per ton to be economically viable as a combustible fuel source. Unfortunately that was not possible. The Japanese have also discovered this to their sorrow, after having spent US$򠄀 million on a 10-year project.

So, do microalgae have a future? Microalgae for food and feed additives have to be produced at a cost of no more than 10 Euros per kg dry mass to be commercially competitive thinks Melkonian, who wonders how the Klötze plant can compete with Japanese facilities that grow the algae faster, heterotrophically—not just using light𠅊nd can sell the product for 30 Euros per kg. As he concluded, ‘I਍oubt they can produce it for this price in Klötze.’ It is a fact that the largest production of unsaturated fatty acids for use as human food additives is accomplished in bacterial fermenters, and heterotrophic algal bioreactors, which are much cheaper than the Klötze method.

Despite the caution of people like Melkonian, there appears to be plenty of venture capital waiting to flow into the green tubes, especially in Germany, and larger companies are also investing in microalgal technology. Microalgae may not be a solution to global pollution after all, they may not even cure disease, but if they help us live longer, healthier and with smoother skin, people will buy them, no matter what the price. As Mende concluded, ‘it’s no wonderdrug, but you sure wonder what it can do next.’

Growing Algae

This project measures the growth rate of algae supplied with supplemental carbon dioxide.

The goal is to have the student conduct a controlled experiment to test a hypothesis about conditions affecting the growth of algae.

  • Does supplemental carbon dioxide affect the growth rate of algae?
  • Is the experimental design capable of producing enough carbon dioxide to drive algal growth?

Algae are organisms commonly found in aquatic environments. There are two types: macroalgae and microalgae. The large multicellular macroalgae are often found in ponds and in the ocean. They tend to be measurable in inches, although giant kelp in the ocean can grow to more than 100 feet in length. Microalgae are tiny unicellular algae that grow as suspensions in water they are measurable in micrometers. Common sources of microalgae are bogs, marshes, and swamps.

All algae require sunlight, water, nutrients, and carbon dioxide for growth. Through the process of photosynthesis, algae convert the carbon dioxide into glucose (a sugar). The glucose is then broken down into fatty acids, which under normal conditions, are used to produce membranes for new algal cells. If, however, the algae are starved of nutrients, the fatty acids produce fat molecules (oil). Because carbon dioxide is the only source of carbon for algae, having an adequate supply is essential if they are to be used for commercial purposes.

  • What materials are required? Three one-liter bottles of purified water sugar brewer&rsquos yeast silicone sealant drill 6-mm aquarium airline tubing algae
  • Materials can be found at the following places: Purified water (supermarket), sugar (supermarket), brewer&rsquos yeast (supermarket), silicone sealant (Walmart-type store) aquarium airline (pet store) algae (pond or marsh or biological/scientific supply house) 10-15-10 liquid plant food (plant nursery or Internet)
  1. Read about the conditions required for algae to grow, and formulate a hypothesis to predict whether giving algae supplemental carbon dioxide would be a feasible way to increase algae growth.
  2. Collect some algae from a pond, marsh, swamp, swimming pool, fish aquarium, bird bath, or other source. If you are unable to locate a natural source, contact a biological/scientific supply house (Google).
  3. Add equivalent amounts of algae to two (clear plastic) bottles of purified water. Discard the bottle caps.
  4. Add two drops of 10-15-10 liquid plant food to each bottle.
  5. Pour out a small amount of water from a third bottle of purified water, leaving about an inch of air space at the top of the bottle. This bottle will be the carbon dioxide reactor.
  6. Make a hole in the bottle cap of the reactor bottle that is just large enough to allow an aquarium airline to pass through it, then run the airline through the hole so that it extends into the free air space when the cap is on. Seal the airline to the top of the bottle cap with a silicone sealant.
  7. Dissolve 2 teaspoons of sugar and 1 teaspoon of brewers yeast in the reactor. (Yeast is a fungus that converts sugar into carbon dioxide bubbles.)
  8. Extend the aquarium airline from the reactor bottle to one of the bottles containing algae. The airline should extend about half way into the algae bottle.
  9. Place all three bottles outdoors where they will get indirect sun. (TIP: Direct sunlight may inhibit growth. The optimum temperature for algal growth is between 20 and 24 degrees C. Temperatures above 35 degrees C are lethal to algae.)
  10. Monitor the growth of the algae in the two sample bottles for one month. If necessary, replace the sugar, yeast, and water in the reactor to keep the carbon dioxide source operating.
  11. At the end of the month, compare the amounts of algae in the two sample bottles.
  12. Evaluate your hypothesis in light of your findings. Revise it if necessary and propose additional experiments.

Terms: Photosynthesis Algae Microalgae Macroalgae Carbon dioxide Yeast Sugar

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A fed-batch strategy is proposed to produce microalgae biomass under non-axenic heterotrophic conditions. The strategy induces the alternation of N-deplete (Glucose-replete) and N-replete (Glucose-deplete) cultivation phases by the periodic and uncoupled supply of glucose and NO3 − to the culture. Cultivation of the microalga T. obliquus with this strategy reduced the ratio of the bacteria to microalgae cell concentration from 1.6, attained by conventional photoautotrophic cultivation, to 0.03. During the N-deplete phase, microalgae duplication stopped and biomass concentration increased 1.9 times, while during the N-replete phase, microalgae duplicated halving their average size and losing about 25% of their weight. The process proved to be effective under several consecutive cycles. Biomass productivity until 6.1 g/Ld and biomass concentration until 26 g/L were achieved. The results demonstrate that the proposed strategy can effectively prevent bacterial contamination, paving the way to the large scale production of microalgae biomass under non-axenic heterotrophic conditions.

Establishment of a bioenergy-focused microalgal culture collection

A promising renewable energy scenario involves growing photosynthetic microalgae as a biofuel feedstock that can be converted into fungible, energy-dense fuels. Microalgae transform the energy in sunlight into a variety of reduced-carbon storage products, including triacylglycerols, which can be readily transformed into diesel fuel surrogates. To develop an economically viable algal biofuel industry, it is important to maximize the production and accumulation of these targeted bioenergy carriers in selected strains. In an effort to identify promising feedstock isolates we developed, evaluated and optimized contemporary high-throughput cell-sorting techniques to establish a collection of microalgae isolated from highly diverse ecosystems near geographic areas that are potential sites for large-scale algal cultivation in the Southwest United States. These efforts resulted in a culture collection containing 360 distinct microalgal strains. We report on the establishment of this collection and some preliminary qualitative screening studies to identify important biofuel phenotypes including neutral lipid accumulation and growth rates. As part of this undertaking we determined suitable cultivation media and evaluated cryopreservation techniques critical for the long-term storage of the microorganisms in this collection. This technique allows for the rapid isolation of extensive strain biodiversity that can be leveraged for the selection of promising bioenergy feedstock strains, as well as for providing fundamental advances in our understanding of fundamental algal biology.


► Bioprospecting protocol established to identify promising microalgal biofuel strains. ► Cryopreservation strategy developed for biodiversity conservation. ► Identification of promising growth media and enrichment strategies.

Green algae

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Green algae, members of the division Chlorophyta, comprising between 9,000 and 12,000 species. The photosynthetic pigments (chlorophylls a and b, carotene, and xanthophyll) are in the same proportions as those in higher plants. The typical green algal cell, which can be motile or nonmotile, has a central vacuole, pigments contained in plastids that vary in shape in different species, and a two-layered cellulose and pectin cell wall. Food is stored as starch in pyrenoids (proteinaceous cores within the plastids). Green algae, variable in size and shape, include single-celled (Chlamydomonas, desmids), colonial (Hydrodictyon, Volvox), filamentous (Spirogyra, Cladophora), and tubular (Actebularia, Caulerpa) forms. Sexual reproduction is common, with gametes that have two or four flagella. Asexual reproduction is by cell division (Protococcus), motile or nonmotile spores (Ulothrix, Oedogonium), and fragmentation.

Most green algae occur in fresh water, usually attached to submerged rocks and wood or as scum on stagnant water there are also terrestrial and marine species. Free-floating microscopic species serve as food and oxygen sources for aquatic organisms. Green algae are also important in the evolutionary study of plants the single-celled Chlamydomonas is considered similar to the ancestral form that probably gave rise to land plants.

Phytoplankton Culture for Aquaculture Feed

This factsheet published by the Southern Regional Aquaculture Center (SRAC) gives information on phytoplankton culture for aquaculture feed.

Phytoplankton consists of one-celled marine and freshwater microalgae and other plant-like organisms. They are used in the production of pharmaceuticals, diet supplements, pigments, and biofuels, and also used as feeds in aquaculture. Phytoplankton are cultured to feed bivalve molluscs (all life stages), the early larval stages of crustaceans, and the zooplankton (e.g., rotifers, copepods) that are used as live food in fish hatcheries.

Flagellates and diatoms are two important types of phytoplankton at the base of the food chain. They manufacture cellular components through the process of photosynthesis, taking up carbon dioxide and nutrients from the water and using light as an energy source.

The microalgae used as feed in hatcheries vary in size, environmental requirements, growth rate, and nutritional value (Fig. 1, Tables 1 and 2) (Helm et al., 2004). When selecting a species for culture, it is important to take all of these parameters into consideration. Most hatcheries grow a variety of species that serve different needs throughout the production cycle with respect to size, digestibility, culture characteristics, and nutritional value (Muller-Feuga et al., 2003).

Culture conditions can vary widely—from outdoor ponds or raceways with nutrients added to promote a bloom of the natural microalgae, to monocultures reared indoors under controlled environmental conditions. This paper focuses on the monoculture of microalgae under clearly defined environmental conditions and production protocols.

Cell Volume, Organic Weight, and Gross Lipid Content of Some Commonly Cultured Phytoplankton Species used in Bivalve Mollusc and Fish Hatcheries (Helm et al., 2004).

Microalgal culture facilities typically use seawater enriched with nutrients—primarily nitrates, phosphates, essential trace elements, vitamins, and, in the case of diatoms, silicates. Water used to culture microalgae should have similar chemical composition to that used to culture the animals, and it should be pretreated. Some laboratories use synthetic seawater for small-scale cultures, but it is prohibitively expensive for large-scale production in commercial hatcheries.

Temperature, Light, and Salinity Ranges for Culturing Selected Microalgae Species (Hoff and Snell, 2008).

Population Dynamics

Algal cells from a starter culture are inoculated into a larger volume of treated, enriched water to reach an initial low density of about 30 to 100 cells/?L. For the first 2 to 3 days the cells become acclimated to the new medium, grow, and begin cell division. This phase, termed the lag phase, varies in length depending on the amount of inoculum used (initial cell density), alga species (inherent division rate), irradiance, and temperature (Fig. 2). Once acclimated, the algal cells divide at an accelerating rate, and the population increases logarithmically this exponential growth phase lasts 4 or more days. The cells are usually harvested for feeding during this phase. The exponential growth phase is followed by the stationary phase, when cell division declines and there is no further increase in cell density. This decreased growth is the result of changes in the concentration of nutrients, self-shading (high cell density reduces the amount of light available to algal cells), and changes in the culture medium, such as increasing pH and the build-up of metabolic waste products or substances called autoinhibitors that are secreted by some species (mostly diatoms). As the culture ages, the stationary phase is followed by a senescent phase in which the density of the culture will decline.

Stationary phase algae should not be used for larviculture because although the algae may be nutritious, as they die the cells rupture and bacteria can proliferate (including some pathogenic bacteria such as Vibrio spp.). The wise culturist knows that the line between feeding larvae and poisoning them can become blurry as algal cultures age.

The Culture Environment

When designing a microalgal production system, consider which species is most appropriate for the intended use (e.g., size and nutritional characteristics). Also consider yield, operating costs, and reliability. Microalgal culture is the most expensive and technically challenging aspect of all hatchery operations. The cost of producing microalgal feed ranges from $100 to $400 per dry kilogram ($45 to $180 per pound) of microalgal biomass (Wikfors, 2000). Algal culture accounts for about 40 percent of the cost of rearing bivalve seed to a shell length of 5 mm in a land-based hatchery (Ukeles, 1980).

Hatcheries use either intensive indoor culture with artificial lighting or extensive outdoor culture in large tanks, raceways, or ponds with natural lighting. Some hatcheries use a combination of the two. Intensive indoor systems are expensive and labor intensive, but they are more reliable and more productive (relative to space requirements) than outdoor systems. Open ponds and raceways are also more prone to biological contamination or other water quality problems. As one might imagine, the potential for “culture crashes” increases as the degree of control over environmental factors such as temperature, illumination, nutrient availability, pH, and potential contamination decreases.

The nutritional value of algae is affected by culture age and growth phase, light characteristics and intensity, nutrient limitation and source, and cell density (Depauw and Persoone, 1988). Whether intensive or extensive, microalgal culture requires filtered and treated water, nutrients, a light source, aeration and mixing, temperature/ salinity control, pH control, and a high-quality inoculum to ensure a satisfactory yield (Fig. 3).

Filtered and Treated Water

Pretreatment of water, whether saltwater or freshwater, is one of the most important steps in successful microalgal culture. Culture water should be free of suspended solids, plankton (e.g., protozoans, ciliates and other algae species), bacteria, unacceptably high concentrations of dissolved organic compounds (DOC), dissolved metals, and pesticides.

Pretreatment typically includes mechanical and chemical filtration, sterilization or disinfection, and nutrient enrichment. The choice of treatment should be based upon species cultured, volume requirements, and cost.

Mechanical filtration. Mechanical filtration removes suspended solids, plankton and bacteria and is usually used with the other forms of treatment described below. The type of mechanical filtration used depends on the condition of the incoming water and the volume of water to be treated. A mechanical filter usually consists of a series of filters that remove increasingly smaller particles—sand filters or polyester filter bags (20- to 35-?m), followed by cartridge filters (10-, 5-, 1-?m) or diatomaceous earth (DE) filters. Small volumes of seawater can be filtered to remove bacteria using 0.22- or 0.45-?m membrane cartridge filters.

Chemical filtration. Dissolved inorganic and organic compounds (DOC), metals, pesticides, and other contaminants can prevent or retard microalgal growth, although detecting them can be complicated and costly. Activated carbon (charcoal) filtration is helpful in reducing DOC, while deionization resins are effective in removing metals and hydrocarbons.

Heat sterilization. Pre-filtered seawater can be sterilized by autoclaving at 1.06 kg/cm2 for 20 minutes. Autoclaving is most suitable for small volumes, while batch or continuous pasteurization at 65 to 70 °C is used for large volumes. Pasteurization at 50 °C for 8 to 10 hours is also effective a glass-lined water heater or 500- to 1,000-W submersion heater can be used. Microwave sterilization is useful for small volumes of pre-filtered seawater (1 to 5 ?m for 8 to 10 minutes per 1 to 1.5 L using a 700-W unit). Nutrients can be added before microwaving since the temperature will not exceed 84 °C (181 °F) (Hoff, 1996). Bellows and Guillard showed that using a 1.2-ft3, 700-W microwave on high power would effectively kill microalgae in 5 minutes, bacteria in 8 minutes, and fungi in 10 minutes in a volume of 1.5 L of filtered (and unfiltered) seawater (Table 3).

Chemical sterilization. Chlorination is the simplest and most common method of chemical sterilization for culture volumes of at least 4 L. Pre-filtered seawater can be sterilized with sodium hypochlorite solution at 2.5 mg/L free chlorine by adding 1 to 5 mL of household bleach (5% sodium hypochlorite) per liter of seawater. Granular swimming pool chlorine is also effective a dosage of 1 ounce (28 g) to 500 gallons (1,875 L) yields a similar concentration of chlorine as liquid bleach. Sterilization occurs in a short period of time, usually 10 to 30 minutes, although many culturists suggest a longer time (12 hours or overnight) for a margin of safety. Before use, neutralize the residual chlorine by adding an excess of sodium thiosulphate solution (Na2S2O3 · 5H2O). If 250 g of sodium thiosulfate is dissolved in 1 liter of water, then 1 mL of the sodium thiosulfate solution added for every 4 mL of bleach used is sufficient to eliminate residual chlorine (dechlorination). Common swimming pool chlorine test kits can be used to determine the presence of residual chlorine, but they do not give a precise measure of chlorine concentration as a general practice, additional sodium thiosulfate solution should be added if there is any indication of residual chlorine.

Summary of Heat Sterilization Types, Effective Methods, Application, and Limitations (Kawachi and Noël, 2005)

Ultraviolet irradiation (UV) and ozone (O3) disinfection. Either UV or ozone can be used to disinfect culture water, although both are most effective after mechanical filtration has removed suspended particulates. It should be noted that “sterilization” is defined as the absolute destruction of all microbial organisms (including bacterial spores), while “disinfection” does not eliminate all microbes but reduces their numbers to a level where the risk of infection is small enough to be acceptable.

UV is the more common of the two, largely because it does not leave concentrations of hazardous by-products. Ozone at high levels can produce chloramines, which are toxic to marine animals. Ozone released into the air can be a safety hazard (if you can smell a faint chlorine smell, residual ozone is present and may be hazardous to your health).

Ozone is a strong oxidizing agent that is particularly effective in removing dissolved organics, pesticides, color and nitrates. It is highly unstable and quickly reverts to O2, but it is also highly corrosive and must be handled with special materials. In-line ozone generators are the most common and usually have monitors/controls to provide an adequate level of ozone yet avoid residual build-up. However, because there is a risk of introducing ozonated water into the culture system, as well as safety concerns for hatchery staff, ozone is not recommended for operators who lack experience and the monitoring equipment to properly manage ozone levels.

Ultraviolet radiation (germicidal energy) is an efficient, simple and reliable way to kill microorganisms in culture water. With proper exposure in clear water, ultraviolet light kills a microorganism by penetrating its cell wall and destroying its nuclear material. Low-pressure mercury vapor UV bulbs are best suited for disinfection because their spectral wavelength (254 nm) is close to the most efficient germicidal wavelength (265 nm). However, the killing power of UV is affected by turbidity/coloration of the incoming water, distance from the source, exposure time (flow rate), species, and age of the bulb (some lights age rapidly, losing as much as 40 percent of their wattage after 6 months). Minimum dosages vary widely for different microorganisms: 15,000 ?watt-sec/cm 2 for most bacteria, 22,000 ?watt-sec/cm 2 for water-borne algae, 35,000 ?watt-sec/cm 2 for bacteria/viruses, 100,000 ?watt-sec/cm 2 for protozoans, and as much as 330,000 ?watt-sec/cm 2 for Aspergillus niger (mold) (Depauw and Persoone, 1988).

Wattage and flow rate are the most important factors in achieving UV sterilization the slower the flow rate, the higher the kill rate for a given bulb wattage (Escobal, 1993). For example, a 40-watt mercury vapor bulb with a flow rate of 500 gallons per hour in a 2-inch-diameter pipe will deliver approximately 11,530 ?watt-sec/cm 2 . Increasing the pipe diameter to 3 inches (thereby reducing flow rate) will increase the dosage to 17,530 ?watt-sec/cm 2 cutting the flow rate in half in that 3-inch pipe further increases the dosage to 34,340 ?watt-sec/cm 2 (Hoff et al., 2008).

Nutrient Enrichment

The objective of culturing microalgae is to obtain the highest cell densities in the shortest period of time, and natural concentrations of nutrients in freshwater and seawater are usually insufficient to support high algal yields. Although trace elements are usually found in sufficient quantities, macronutrients are in short supply (usually phosphorus in freshwater and nitrate in saltwater). Several nutrient enrichment media containing soil extract, nitrates, phosphorus, trace elements, and vitamins have been described for freshwater and saltwater (Creswell, 1993). Of the nutrient media formulations used to culture marine microalgae in laboratories and hatcheries, Guillard and Ryther’s F/2 media is the most widely used, and a pre-mixed solution is available from a variety of vendors (Table 4). There are dozens of culture media recipes, many of which were formulated specifically for certain types/species of microalgae and cyanobacteria. A good reference is Algal Culturing Techniques, edited by R. A. Anderson. Table 5 lists services that have culture medium recipes on their websites.

Guillard’s F/2 Media used to Culture Marine Microalgae (Guillard, 1975).

Major Service Culture Collections with Culture Medium Recipes on Their Websites (Anderson, 2005).

Light Source

Light is the energy source that drives photosynthesis to convert nutrients into algal biomass. Maximum culture depth and cell density are the primary variables regulating the efficient use of light (Richmond et al., 1980). Light intensity, spectral characteristics, and photoperiod are the components of an illumination regime. Indoor microalgal facilities usually use fluorescent “cool white” bulbs (2,500 lux), while outdoor systems and greenhouses use ambient sunlight in combination with fluorescent or metal halide bulbs to provide evening illumination. The spectral characteristics of “cool white” bulbs are not ideal for intensive microalgae production bulbs with enhanced red and blue wavelengths (Gro-LuxTM) support higher yields. The age of the bulb is also important, as the spectral characteristics and luminosity change over time bulbs should be replaced at least annually.

Irradiation of 2,500 to 5,000 lux (250 to 500 footcandles) is optimal for microalgae photosynthesis, with a maximum of 10,000 lux (Escobal, 1993). Guillard (1975) recommended 3,500 and 4,500 lux for stock culture of Thalassiosira pseudonana under continuous and 14 hours per day illumination, respectively. In indoor facilities, bulbs should be 6 to 10 inches from stock cultures if possible, the ballasts should be outside the culture room to help maintain temperature control.

Internally illuminated culture vessels are costly to construct but inexpensive to operate. Mounting the lamps inside a glass or clear plastic cylinder within the culture vessel reduces the distance the light must travel to penetrate the culture. Culture cylinders with internally mounted lights typically produce as much as cultures with three times the volume.

Metal halide lamps (750 to 1,000 W) are usually used to illuminate larger cultures (1,800 liters or more), and since they generate considerable heat, they should be placed at least 12 inches above the surface in open, well-ventilated greenhouses. If natural light is being used for large-volume cultures in greenhouses, it is best to use the morning sun, with 40 percent shade cloth on the west side of the building and 60 percent shade cloth down the middle of a northsouth oriented building, especially in summer.

Although most commercial light meters measure “lux,” many references in the literature related to light requirements for phytoplankton culture prefer to express optimum irradiance in terms of “Photosynthetically Active Radiation” (PAR), which is expressed as ?mol photons · s -1 · m -2 , radiation within wavelengths of 400 to 700 nm. Converting lux values to PAR depends on the type of lamp and its spectral characteristics. Multiply lux by the following conversions for PAR:

  • incandescent = 0.019
  • metal halide = 0.014
  • cool white flourescent = 0.013
  • daylight flourescent = 0.014
  • GRO flourescent = 0.029
  • clear day sunlight = 0.018

Artificial light is usually preferred over sunlight. With sunlight, the duration and intensity are not easily controlled, which may cause overheating, insufficient irradiance, or photoinhibition if the light is too intense for too long (Escobal, 1993). Artificial lighting can be controlled with a simple timer or light monitor and should be set for a minimum of 16:8 hours light/dark per day (minimum) to 24-hour illumination (maximum) for indoor cultures. Although artificial lighting can be precisely controlled in terms of quality and quantity, it is costly and accounts for almost 95 percent of the cost to culture microalgae (Muller-Feuga et al., 2003).

Temperature Control

Because most of the microalgae species preferred by culturists are tropical/subtropical, most strains grow best at temperatures ranging from 16 to 27 °C (60 to 80 °F). The optimum is about 24 °C (75 °F). Ukeles (1976) compared the growth response of several microalgae species to temperature (Table 6). The optimum temperature for growth will vary with species, and to some extent is a complex factor that depends on other environmental conditions. Cultures should be maintained at the lowest temperature that is consistent with good yield to avoid encouraging bacterial growth. When considering temperature characteristics for an enclosed culture room, one should consider: 1) the size of the room, 2) heat sources (such as lights and ballasts), and 3) the volume and temperature of air pumped into the culture vessels.

Growth Response of Different Microalgal Species to Various Temperatures (°C). Growth Rates are Relative to Performance of a Control Cultured at 20.5 °C (Ukeles, 1976).

Aeration and Mixing

Aeration is important for microalgal culture because: 1) air is a source of carbon (from CO2) for photosynthesis 2) CO2 provides essential pH stabilization and 3) physically mixing the culture keeps nutrients and cells evenly distributed, reduces self-shading and/or photoinhibition (a decrease in photosynthesis due to excess light), and avoids thermal stratification in outdoor systems. Air diffusers (airstones) create small bubbles that maximize oxygen/ CO2 transfer, and they are frequently used for smallvolume cultures. In larger culture containers, fine bubbles from air diffusers create spray and foam that can promote bacterial growth larger bubbles (no airstones) actually do a better job of mixing the culture with minimal foaming. Common alternatives for mixing larger volume cultures include jet pumps, paddle wheels, continuous recirculation, and air-lift pumps (Persoone et al., 1980).

Carbon dioxide (CO2) source and pH control

Carbon dioxide plays a dual role in microalgal culture. It provides a source of carbon to support photosynthesis, and it helps maintain pH at optimum levels (7.5 to 8.2 for marine species). As culture density increases, more carbon is consumed through photosynthesis, reducing CO2 concentration and causing the pH to rise. At about pH 10 some nutrients will precipitate, algal growth will be retarded, and the culture could completely collapse. This can be prevented if the pH is maintained by introducing CO2 into the air delivery system. This can be done manually (while the cultures are illuminated), pulsed intermittently using a timer and solenoid valve, or, most effectively, by using a pH monitor/controller.


Most hatcheries will culture several species of microalgae to provide live feeds with different sizes and nutritional characteristics, depending on the animal being cultured and its life stage. The culture protocol for each species will be dictated by the characteristics of the microalgae (e.g., growth rate and environmental requirements), harvest yields, and use requirements.

Maintaining and Transferring Stock and Starter Cultures

Stocks of monospecific (uni-algal) cultures can be obtained by collecting local species, separating them by size (filtration) or density (centrifugation), and inoculating agar plates containing enrichment media. From these multi-species algal cultures, individual colonies are selected through agar streaks, micropipette isolation, liquid dilution, or flow cytometer cell sorters (Fulks and Main, 1991). Culturing algae in highly filtered, autoclaved, enriched seawater in the presence of antibiotics allows bacteria- and protozoanfree pure cultures (axenic) to be isolated. However, the isolation and screening of local species is laborious and mono-specific cultures of most microalgae species used for aquaculture are readily available from research laboratories, commercial hatcheries, and vendors.

Stock cultures are kept in specialized maintenance media, which may be enriched seawater or nutrientenriched agar plates or slants, under closely controlled conditions of temperature and illumination. A special temperature-controlled area or room adjacent to the algal culture room is usually allocated for this purpose.

Stock cultures serve as inocula for the large-volume production of phytoplankton used for harvest or feeding. Stock cultures containing sterile, autoclaved media are kept in small, transparent, autoclavable containers such as 25-mL test tubes or 250- to 500-mL borosilicate glass, flat-bottomed, conical flasks fitted with cotton wool plugs at the necks (or polyethylene beakers can serve as caps). They also are maintained in seawater agar medium impregnated with suitable nutrients in petri dishes or on slants in test tubes. Every effort should be made not to contaminate the stock and starter cultures with competing microorganisms. To minimize potential contamination, an enclosed culture transfer hood outfitted with a Bunsen burner and UV lights should be used (a laminarflow hood is preferred if available) (Fig. 4).

The sterile procedures described below should be followed.

  1. Wipe all inner surfaces of the inoculating hood and working surfaces with 70 percent ethanol.
  2. Place all flasks that will be used in the hood, including flasks to be transferred from (the transfer flask) and flasks containing sterilized media that will be inoculated under the culture transfer hood.
  3. Irradiate flasks to be inoculated with an ultraviolet lamp for at least 20 minutes. Be sure the hood has a dark cover over the viewing glass (UV radiation can be damaging to the eyes).
  4. Switch off the UV lamp ignite a small Bunsen burner remove caps from one transfer and one new flask and flame the neck of each flask by slowly rotating the neck through the flame.
  5. Tilt the neck of the transfer flask toward the new flask. In one motion, remove both stoppers and pour an inoculum into the new flask. Transfer approximately 50 mL for diatom species and 100 mL for flagellates. Avoid touching the necks of the two flasks. Never touch the portion of the stopper that is inserted into the flask. Once the inoculum is added, replace the stopper in the transfer flask. Slowly flame the neck of the new flask before replacing its stopper.
  6. Replace the cap over the neck of the new flask and use a waterproof marker pen to label the new flask with the algal species inoculated and the date of transfer.
  7. After all inoculations are completed, turn off the burner and transfer all new flasks to an algal incubator or a well-lit area in the algal culture facility. The remaining inoculum in the transfer flasks can be used to inoculate larger cultures such as 4-L flasks or carboys.
  8. Empty test tubes, flasks, stoppers and/or caps should be removed, thoroughly washed, and sterilized or discarded.
  9. Remove all materials from the working area and wipe the surface with 70 percent ethanol.

If you are transferring liquid cultures using glass Pasteur pipettes, follow these steps (Kawachi and Noel, 2005):

  1. Bring the pipette canister (used to sterilize the pipettes) close to the Bunsen burner, the cap removed, and gently shake it so that one pipette is extruded a few centimeters from the canister opening.
  2. Remove the pipette from the canister carefully so that its tip does not come in contact with the canister opening.
  3. Replace the canister lid. Place the pipette bulb adjacent to the Bunsen burner and clean the inside with 70 percent ethanol.
  4. Place the bulb on the pipette, pick up the cell culture vessel, and flame at an angle of at least 45 degrees.
  5. Remove the vessel from the flame, insert the tip of the pipette into the liquid, being careful not to touch the sides of the vessel, and collect the desired amount of inoculum by controlling the pressure of the bulb.
  6. Remove the pipette, orienting it in an almost horizontal position reflame the mouth of the vessel, and replace the cap or plug.
  7. Using the same procedure, open the new vessel, flame the opening, and insert the pipette into the new vessel without touching the mouth.
  8. Slowly discharge the cell suspension, remove the pipette, flame the mouth of the vessel, and replace the cap.
  9. Remove the pipette bulb and place the used pipette into a discard container to be discarded or reused. Clean the bulb with 70 percent ethanol for reuse.
  10. Once all transfers are completed, turn off the Bunsen burner, remove all materials from the working area, and wipe the surface with 70 percent ethanol.

Progressive Batch Culture

The quantities of algae cells required for feeding mollusc larvae and other zooplankton are produced through a process called progressive batch culture (transferring small-volume cultures of concentrated inoculum into larger volumes of treated, enriched water). Starting with cells taken from an axenic stock culture (test tubes), microalgae are cultured in an enriched medium through a series of culture vessels of increasing volume (Fig. 5). The algae grown in each culture vessel serves as the inoculum for the next larger vessel, until the quantity of cells required for feeding is reached. This is a typical series for large-scale production:

  1. 25-mL test tubes (10-mL stock culture) inoculates….
  2. 500-mL flasks (250-mL starter culture) inoculates….
  3. 2.8- to 4-liter flasks (1,000-mL culture) inoculates….
  4. 20-liter carboys (16-liter culture) inoculates….
  5. 250-liter cylinders (180-liter culture) inoculates….
  6. 12,000-liter tanks (10,000-liter culture) inoculates….

Starter cultures are used to inoculate “intermediate cultures” (2 to 25 L), which are used to inoculate even larger volume cultures for final production before harvest and feeding. Similar to stock cultures, starter cultures can be grown in 500-mL flasks with 250 mL of sterile medium about 50 mL of the starter culture is transferred to similar volume flasks to maintain the line, while the remaining 200 mL are used to inoculate intermediate culture containers (typically from 4-L flasks to 20-L carboys) (Fig. 6).

Procedures for maintaining starter cultures are almost identical to those described above for stock cultures. A line of starter cultures is originally established from the stock culture of the required species. They are grown at 18 to 22 °C at a distance of 15 to 20 cm from 65- or 80-W fluorescent lamps, giving a level of illumination at the culture surface of 4,750 to 5,250 1ux. Small-volume cultures (test tubes to 1-L flasks) are usually manually shaken daily to facilitate gas exchange and mixing. Aeration is required for 2-L flasks and larger volumes, and in-line filters on the delivery tubing are necessary to prevent contaminants that can be introduced through aeration. Starter cultures are generally aerated with an air/CO2 mixture to maintain a satisfactory pH and provide additional carbon for photosynthesis. When CO2 is used, the pH is usually maintained between 7.5 and 8.5.

Stock cultures are grown for varying periods before inoculation in 500-mL flasks. For diatom species this period is 3 to 5 days for the majority of flagellates it is 7 to 14 days. When ready for use, 20 to 50 mL of the starter culture (depending on species and cell density) is transferred to a fresh 250-mL culture to maintain the starter culture line. The remainder is used as inoculum for larger cultures (usually 1,000 mL in 2.8- to 4-L flasks) to be grown for feeding or as an intermediate step in large-scale culture, where they are used as inocula for 20- to 40-L cultures. To maintain high-quality cultures, transfers should be made during the exponential growth phase, with an inoculum of at least 10 to 20 percent of the total volume or an initial concentration of about 105 cells/mL, to promote rapid population growth.

Throughout the scale-up process, contamination is a constant threat and cleanliness and attention to detail are critical. Contaminants may be chemical or biological and they can originate from one or several sources. A common chemical contaminant is residual chlorine from the sterilization process, while biological contaminants might include: 1) excessive levels of bacteria (indicated by cloudy water), 2) protozoans or rotifers (culture water turns off color and clears), 3) competing microalgae (color change or crust attached to culture vessel walls), and 4) macroalgae (green or brown strands attached to culture vessel walls). Identifying bacterial and microalgal contaminants usually requires 100X to 400X magnification, while protozoan contamination can be observed under 15X to 40X magnification (Hoff and Snell, 2008). Possible sources of contamination are shown in Figure 7.

Estimating Algal Density

Estimating algal density is an inherent part of any algal production system. Algal biomass is the criterion used to determine when to transfer inoculum through serial dilution to larger volume cultures and to determine harvest volumes of cultures in production. For stock and starter cultures, the most accurate measurement of cell density can be made using a Palmer-Maloney slide or a hemacytometer.

The chamber of the Palmer-Maloney slide is without rulings and is circular. It is 17.9 mm in diameter, 400 ?m deep, and has an area of 250 mm 2 , for a total volume of 0.1 mL. Even very small microalgae at low concentrations (10 per mL) can be detected. Hemacytometers are thick glass slides with two chambers on the upper surface, each measuring 1.0 x 1.0 mm. A special coverslip is placed over these two chambers, giving a depth of 0.1 mm and a total volume in each chamber of 0.1 mm 3 . The base of each chamber is marked with a grid to aid in counting cells within the area (Fig. 8). Before counting motile algal species, one or two drops of 10 percent formalin should be added to a 10- to 20-mL sample of the culture to be counted. With the coverslip in position, one or two drops of the algal sample are introduced by means of a Pasteur pipette to fill both chambers.

The central grid of each chamber is subdivided into 25 squares, each measuring 0.2 x 0.2 mm these are further subdivided into 16 squares (0.05 x 0.05 mm). Therefore, the volume of each grid is 0.2 x 0.2 x 0.1 mm = 0.004 mm 3 . To determine cell density:

  • Count the number of cells in ten randomly selected 0.2 x 0.2 mm grids and calculate the average (as an example, an average of 42.5 cells/grid).
  • Multiply the average (42.5 cells) by 250 to get 10,625 cells/mm 3 (0.004 mm 3 x 250 = 1 mm 3 ).
  • Since there are 1,000 mm 3 in 1 mL, multiply the value in the second step by 1,000 to get cells/mL. 42.5 x 250 x 1,000 = 10,625,000 cells/mL, 10.625 million cells/mL (10.62 x 10 6 ).

The Coulter Counter, now called a “multisizer,” was originally developed to count blood cells. Algal cells pass through a small aperture (2 to 10 ?m) and a slight electrical current travelling between two electrodes. Each time a cell passes between the electrodes, the current is impeded and the cell is counted. The advantages of the Coulter Counter are its accuracy and efficiency the disadvantages are that it does not discriminate between algal cells and other particles, dense culture needs to be diluted to get an accurate count, and they are expensive.

For larger cultures, a spectrophotometer or fluorometer that measures the chlorophyll ? content in an algal culture can be used to obtain a quick approximation of cell density. Graphs comparing cell density and readings on either instrument must be prepared for each algal species. However, the chlorophyll ? content in an algal cell is not constant and varies with the nutritional state of the cell. This will affect the accuracy of cell density estimates derived with these instruments.

An inexpensive way to estimate algal density in large cultures is to use a “Secchi disk,” a technique that has been used by field biologists for decades. Once calibrated to the microalga species in culture, Secchi disks can provide a reasonably accurate estimate of algal cell density (Hoff and Snell, 2008).

Intermediate Culture

Intermediate culture volumes, typically 4-L flasks to 20-L carboys, are used to inoculate larger vessels, typically 100- to 200-L translucent fiberglass cylinders or polyethylene bags, or even larger fiberglass tanks and raceways. The complexity of the culture operation depends on the requirement for algae and cost constraints. The simplest culture system may be just a scaled-up version of the starter cultures using 4-L flasks or 20-L carboys. Sterile, nutrient-enriched seawater with an inoculum should be aerated with a mixture of 2 percent CO2 carried in compressed air. Illumination for culture growth is provided by fluorescent lamps, usually mounted externally to the culture flasks. The number of lamps used is determined by the height and diameter of the culture vessels, with the object of providing 15,000 to 25,000 lux measured at the center of the empty culture container. Two 65- or 80-W lamps are sufficient to illuminate 3-L glass flasks, which are about 18 cm in diameter, whereas five lamps of the same light output are necessary for 20-L carboys (Fig. 9).

Cultures 4 to 8 days old from carboys (20-L) are used to inoculate 200-L translucent fiberglass cylinders or polyethylene culture bags. In most cases, these larger volume cultures are housed in greenhouses and receive natural light (adequate illumination from fluorescent bulbs is usually cost prohibitive). The vessels are filled with filtered, UV-irradiated, and usually chlorinated/dechlorinated seawater, enriched and inoculated. Each cylinder is carefully labeled to document the date of the sterilization process, the enrichment, and the species inoculated. Under optimal environmental conditions, the culture will be harvested in 4 days or used to inoculate large-scale cultures in tanks or outdoor ponds. (Fig. 10).

Batch, Semi-continuous, and Continuous Culture Systems

Because microalgal culture produces high concentrations of cells, most laboratories require only small volumes of algae for food. These can be cultured in 4-L containers or 20-L carboys using “batch” culture protocols. Commercial hatcheries, which require much larger volumes of algae, often use semi-continuous or continuous culture systems.

Batch cultures are inoculated with the desired species that will grow rapidly under optimal conditions until the rate of cell division begins to decline, indicating the transition from the exponential phase to the stationary phase. At that point, the culture is completely harvested and the container is washed, refilled (with sterilized, enriched medium), and inoculated to begin a new culture. Batch culture is generally used for delicate species or for rapidly growing diatoms. Although batch culture is considered the least efficient method of production, it is predictable, and contamination is less likely than in semi-continuous cultures that stay in production through several harvests. Because the entire culture is harvested, yield per tank is less than in semi-continuous systems therefore, more tanks are required for the same level of production (Fulks and Main, 1991).

Semi-continuous cultures begin much the same way as the batch cultures, but instead of harvesting the entire volume, 25 to 50 percent of the volume is harvested at the point when light has become a limiting factor (late exponential phase). The harvested volume is then replaced with freshly prepared culture medium and the remaining algal cells serve as inoculum. Semi-continuous cultures grow rapidly and can be harvested every 2 or 3 days. In this way, the life of a culture can be extended cultures of some hardier species, such as Tetraselmis suecica, will last for 3 months or more with harvests of 25 to 50 percent of the culture volume three times each week. Semi-continuous culture is mainly used with hardier species of flagellates. Semi-continuous cultures may be grown indoors or outdoors. Their longevity is unpredictable, especially outdoors, because competitors, predators, bacteria and/or other contaminants and metabolites build up and render the culture unsuitable (Guillard and Morton, 2003).

Droop (1975) defines continuous culture as “steadystate continuous flow cultures in which the rate of growth is governed by the rate of supply of the limiting nutrient.” Continuous culture systems are delicately balanced so that culture organisms are harvested continually and the nutrient-enriched media is replenished continually, consistent with the growth rate (sustainable yield) of the culture.

In order to harvest algae continuously at a level adjusted to the maximum specific growth rate (exponential phase) of the culture species, two monitoring and control devices can be used—chemostats and turbidostats. In both cases, fresh, sterile media enters the culture vessel, displacing old media and algal cells that are harvested through an overflow port. Chemostats act on the principle of limited nutrients, so if the concentration of the limiting nutrient (e.g., nitrate) falls below a certain level, a fixed quantity of nutrient solution is added algal growth rate is regulated by the limiting nutrient, not cell density, and flow is continuous (James et al., 1988). Turbidostats have photoelectric monitors connected to solenoid valves that control the withdrawal of algal suspension and the addition of fresh medium as a function of cell density (by measuring turbidity) the flow is not continuous. A variety of mathematical models have been developed that, theoretically, can maximize production from continuously harvested systems (based on algae growth rate, optimum dilution rate, and nutrient concentration), but in practice the culture manipulations are determined empirically after a series of several trials (Sorgeloos et al., 1976 Laing and Jones, 1988 Landau, 1992).

Culture in Polyethylene Bags

Heavy-gauge polyethylene tubing can be cut to a suitable length and one end heat-sealed to form a sterile, flexible culture container that is either a cylinder or an oblong bag (Baynes et al., 1979 Trotta, 1981). The culture vessel design is based on that used by SeaSalter Shellfish Company Ltd. (Farrar, 1975). Containers formed in this way can be strengthened by supporting them within a plasticcoated, steel-mesh frame (Fig. 11). Or, the cylinders can be suspended, with or without lateral support mesh, if the diameter of the bag is less than 30 cm and the height less than 200 cm.

Continuous microalgal culture in polyethylene bags has several advantages: 1) a sealed container is less likely to become contaminated than a rigid container with an open top or lid 2) bags do not require daily maintenance and cleaning and 3) they cost less to install and use space more efficiently. Bags are the least expensive way of constructing large-scale culture vessels. Such containers can be used indoors with artificial illumination or outdoors in natural light. Polyethylene bags have a relatively short lifespan because the internal surface attracts culture debris and bacteria that reduce light penetration and are a source of contamination. At the end of a culture run, it is necessary to replace the bag.

Large-scale outdoor bag cultures are often positioned horizontally to maximize sunlight penetration (Fig. 12). Such large-volume systems are often used to induce multi-species blooms that are best suited for feeding juvenile shellfish in nursery systems or adult shellfish in broodstock systems, rather than for hatchery production. The rate at which a bloom develops is related to the species composition the volume and cell density of the inoculum the quantity, quality and duration of light nutrient levels and temperature.

Concentrating Algal Biomass

In most hatcheries, microalgae are fed in liquid form directly to the animal culture tanks. Recently, though, there has been an interest in concentrating algae to reduce the volume of microalgal culture water (and possible contaminants) introduced into culture tanks. The use of “algal paste” or concentrate has gained popularity because during periods of excess production, the concentrate greatly reduces the physical space required, can be refrigerated until needed, and can be diluted when used. However, the nutritional quality of the microalgae may be a concern if the concentrate is stored for extended periods. Although this concept is not new to aquaculture (Barnabe, 1990), algal paste in preserved or fresh forms recently has become commercially available.

For small-scale harvests, filter screens or cartridge filters (1 to 5 ?m) are effective. The concentrated cells are washed off with limited water and then used, refrigerated or preserved (Hoff and Snell, 2008). Chemical flocculation using natural organic agents such as gelatin, chitosan, and sodium alginate can be used to concentrate microalgae to feed benthic detritus feeders and crustaceans. Centrifugation is used to concentrate large-volume cultures. Continuous- flow centrifuges (e.g., Sharples PenwaltTM) are used to concentrate microalgal cultures into paste. Both chemical flocculation and centrifugation have proven suitable in terms of efficiency and cell density when preparing concentrations for aquaculture feeds (Heasmann et al., 2000).

Advanced Algal Production Systems

There are several new microalgal production systems on the market they are collectively termed “AAPs” (Advanced Algal Production Systems) or photobioreactors. Photobioreactor systems can provide higher algal densities, more efficient space usage (a smaller footprint), continuous or semi-continuous production, longer production cycles with less contamination, and lower labor requirements (Ellis and Laidley, 2006). In general, three types of systems are under production: 1) tubular photobioreactors, 2) column or cylinder photobioreactors, and 3) flat-panel or plate photobioreactors (Tredici et al., 2009).

Closed photobioreactor systems have several advantages over conventional tank-based or pond microalgal productions systems. Still, they have limitations, such as overheating, oxygen accumulation, biofouling, and shear stress (Ellis and Laidley, 2006). Many of these designs are still under evaluation, so the reader is advised to review the specifications of these systems, including purchase and construction costs, operational efficiency, practical application to your production needs, etc. (Anderson, 2005 Tredici, et al., 2009).

Second Generation Biofuels: High-Efficiency Microalgae for Biodiesel Production

The use of fossil fuels is now widely accepted as unsustainable due to depleting resources and the accumulation of greenhouse gases in the environment that have already exceeded the “dangerously high” threshold of 450 ppm CO2-e. To achieve environmental and economic sustainability, fuel production processes are required that are not only renewable, but also capable of sequestering atmospheric CO2. Currently, nearly all renewable energy sources (e.g. hydroelectric, solar, wind, tidal, geothermal) target the electricity market, while fuels make up a much larger share of the global energy demand (∼66%). Biofuels are therefore rapidly being developed. Second generation microalgal systems have the advantage that they can produce a wide range of feedstocks for the production of biodiesel, bioethanol, biomethane and biohydrogen. Biodiesel is currently produced from oil synthesized by conventional fuel crops that harvest the sun’s energy and store it as chemical energy. This presents a route for renewable and carbon-neutral fuel production. However, current supplies from oil crops and animal fats account for only approximately 0.3% of the current demand for transport fuels. Increasing biofuel production on arable land could have severe consequences for global food supply. In contrast, producing biodiesel from algae is widely regarded as one of the most efficient ways of generating biofuels and also appears to represent the only current renewable source of oil that could meet the global demand for transport fuels. The main advantages of second generation microalgal systems are that they: (1) Have a higher photon conversion efficiency (as evidenced by increased biomass yields per hectare): (2) Can be harvested batch-wise nearly all-year-round, providing a reliable and continuous supply of oil: (3) Can utilize salt and waste water streams, thereby greatly reducing freshwater use: (4) Can couple CO2-neutral fuel production with CO2 sequestration: (5) Produce non-toxic and highly biodegradable biofuels. Current limitations exist mainly in the harvesting process and in the supply of CO2 for high efficiency production. This review provides a brief overview of second generation biodiesel production systems using microalgae.

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Build Your Own Photobioreactor

This project attempts to come up with a design for a growing chamber that will meet the physiological requirements of algae for maximum growth.

The goal is to give the student an opportunity to use scientific methodology to design test a photobioreactor design.

  • Can an inexpensive photobioreactor design be found to accommodate the growth needs of algae?
  • What kinds of light sources will optimize the growth of algae?
  • Can a suitable pH be maintained in the photobioreactor?
  • Can a mixing scheme be developed to stir the algae?
  • Can a suitable temperature be maintained in the reactor?
  • Can a suitable nutrient source be found for the algae?
  • Can the initial photobioreactor design be improved?

Algae require sunlight, a source of carbon, nutrients (nitrogen or silicon), and water for growth. They also benefit from stirring.

Algae require light energy in order to convert carbon dioxide into the organic compounds required for growth. If exposed to direct sunlight, their growth may be inhibited. If artificial lighting is used, fluorescent tubes emitting blue or red light are preferred. Artificial light should be available for 18 hours a day.

The pH of the water in which algae are grown should be between 7 and 9.

Stirring is desirable so that all the algae are equally exposed to light and nutrients. The algae should be stirred daily. Not all algae can tolerate vigorous mixing, however.

The temperature of the water should be between 20 and 24 deg C. Temperatures below 16 deg C will slow down growth, while temperatures above 35 deg C are lethal to algae.

A photobioreactor is a chamber that houses and cultivates algae. It maintains suitable conditions of light, nutrients, air, and temperature for the culture.

  • The materials required to complete this project are entirely up to the student. A very basic photobioreactor might make use of three one-liter bottles of purified water sugar brewer&rsquos yeast silicone sealant drill 6-mm aquarium airline tubing and algae (see the Science Fair Project entitled &ldquoCarbon Dioxide and Algae&rdquo and submitted in this group.)
  1. Review the growth requirements of algae.
  2. Design an inexpensive photobioreactor that will cause the algae to grow faster. Keep in mind the following:
  • The light should not be too intense. If algae are exposed to direct sunlight or if they are too close to an artificial light, their growth may be inhibited. If you opt for artificial light, you should use a fluorescent tube that emits either blue or red light. (These are the most active parts of the light spectrum for photosynthesis.) The light will need to be on at least 18 hours each day.
  • An algae culture can completely collapse if the pH falls out of the range between 7 and 9. During algal growth, pH may climb to 9, which may be counterbalanced by addition of carbon dioxide to the reactor.
  • Mixing the algae ensures that all of the algae cells are exposed to the light and nutrients. Mixing also prevents temperature gradients from forming in outdoor bioreactors.
  • Most strains of algae prefer a temperature range between 20 and 24 deg C, although temperatures between 16 to 27 deg C are usually tolerated. Many algae die at temperatures above 35 deg C.
  • Algae require nutrients for optimal growth.
  1. Collect some algae from a natural source such as a pond, marsh, swamp, swimming pool or bird bath. If you are unable to locate a natural source, try a biological/scientific supply house.
  2. Measure the amount of algae collected.
  3. Introduce the algae into the photobioreactor.
  4. Measure the growth of the algae after two weeks. Modify the design of the photobioreactor as needed with an eye toward improving algae yield.

Terms: Photosynthesis Algae Carbon dioxide Light pH Nutrients Photobioreactor

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Integration of microalgae cultivation with industrial waste remediation for biofuel and bioenergy production: opportunities and limitations

There is currently a renewed interest in developing microalgae as a source of renewable energy and fuel. Microalgae hold great potential as a source of biomass for the production of energy and fungible liquid transportation fuels. However, the technologies required for large-scale cultivation, processing, and conversion of microalgal biomass to energy products are underdeveloped. Microalgae offer several advantages over traditional ‘first-generation’ biofuels crops like corn: these include superior biomass productivity, the ability to grow on poor-quality land unsuitable for agriculture, and the potential for sustainable growth by extracting macro- and micronutrients from wastewater and industrial flue-stack emissions. Integrating microalgal cultivation with municipal wastewater treatment and industrial CO2 emissions from coal-fired power plants is a potential strategy to produce large quantities of biomass, and represents an opportunity to develop, test, and optimize the necessary technologies to make microalgal biofuels more cost-effective and efficient. However, many constraints on the eventual deployment of this technology must be taken into consideration and mitigating strategies developed before large scale microalgal cultivation can become a reality. As a strategy for CO2 biomitigation from industrial point source emitters, microalgal cultivation can be limited by the availability of land, light, and other nutrients like N and P. Effective removal of N and P from municipal wastewater is limited by the processing capacity of available microalgal cultivation systems. Strategies to mitigate against the constraints are discussed.

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