Collecting freshwater algae
Macroalgae and the attached microalgae can be collected by hand or with a knife, including part or all of the substrate (rock, plant, wood etc.) if possible. Search all habitats in the waterbody including the edge of stones in fast-flowing water, aquatic plants, dam walls, and any floating debris.
In running or slightly turbid waters, a simple viewing box made from transparent perspex enables attached algae to be more easily observed. A hand lens is often useful to determine if material is reproductive (essential for species determination in some genera and helpful for generic placement).
Microscopic floating algae (the phytoplankton) can be collected with a mesh net (e.g. with 25-30 µm pores) or, if in sufficient quantity (i.e. colouring the water), by simply scooping a jar through the water. Water samples can be left overnight allowing the algae to settle and concentrate on the bottom of the container. Squeezing Sphagnum and other mosses, or some aquatic flowering plants such as Utricularia is a good way to collect a large number of species.
Microscope slides suspended in a waterbody for c. 2-4 weeks will reveal many species. The slides should be kept submerged until ready to examine under the microscope. One side can be wiped clean and a coverslip placed over the other.
Algae growing on soil are difficult to collect and study, many requiring culturing before sufficient and suitable material is available for identification.
The collector's name and a collecting number should be pencilled onto the herbarium sheet (and later replaced with a full label) or onto a collecting tag inserted into the solution. The accompanying notes should include standard information such as the locality, date of collection, and the collector's name and collection number, and as many of the following features as possible: whether the water is saline, brackish or fresh; whether the collection site is terrestrial, or a river, stream or lake; whether the alga is submerged during water level fluctuations or floods; whether the water is muddy or polluted; whether the alga is free floating or attached, and if the latter, the type of substrate to which it is attached; and the colour, texture and size of the alga.
Examining freshwater algae
Microscopes & microscopy
Examining fresh material
Observations (preferably including drawings or photographs) based on living material are essential for the identification of some genera and a valuable adjunct to more leisurely observations on preserved material for others. The simplest method is to place of drop of the water including the alga onto a microscope slide and carefully lower a coverslip onto it. It is always tempting to put a large amount of the alga onto the slide but smaller fragments are much easier to view under a microscope. Start by observing the algae at lower magnification (X40 or X100) and move sequentially up if necessary. Microalgae may be better observed using the 'hanging drop method': place a few drops of the sample liquid on a coverslip and turn it over onto a ring of paraffin wax, liquid paraffin or a 'slide ring'.
A permanent slide is a valuable addition to wet and dry herbarium specimens. Analine blue (1% aqueous solution with 4% molar HCl), Toluidine blue O (0.05% aqueous solution) and Potassium permangenate (2% aqueous KMnO4) are useful stains for macroalgae (different stains suit different species) and Indian Ink is a good stain for highlighting mucilage and some flagella-like structures.
After staining for 30 seconds to five minutes (depending on the material), rinse in water, then add a drop or two of 10% corn syrup solution (Karo™ corn syrup with the addition of 2% phenol) to a small piece of the algae placed on a microscope slide then carefully lower the coverslip. (A corn syrup solution of 5% or less may be required for the more fragile species.) Add drops of 40% corn syrup solution at the side of the coverslip as the liquid underneath the coverslip evaporates. Once ‘set’ (i.e. solid but often still sticky), the sides of the coverslip can be readily sealed with nail polish.
Glycerine solution (75% glycerine, 25 % water) is another useful mounting agent and should be introduced in a similar manner to the corn syrup, i.e. starting with a very dilute solution and building up to 100% glycerine. Sealing with nail polish is essential.
These two mountants are unsuitable for most unicellular algae which should be examined fresh or in temporary mounts of liquid-preserved material.
Magnifications of between 40 and 1000 times are required for the identification of all but a few algal genera. A compound microscope is therefore an essential piece of equipment for anyone wishing to discover the world of algal diversity. Student microscopes with 10X eyepiece and 4X-10X-40X objectives are available for $410-$520 and such a microscope would be suitable for identifying all algae in this guide. An oil immersion 100X objective, available for c. $90, would be a useful addition, particularly when identifying to species level. A camera lucida attachment is helpful for producing accurate drawings while an eyepiece micrometer is important for any species-level identifications. Phase-contrast or interference (e.g. Nomarski) microscopy can improve the contrast for bleached or small specimens, but are available only on more expensive microscope systems.
A dissecting microscope providing magnifications up to 40 or 50 times is a useful aid but is secondary to a compound microscope. High quality dissecting microscopes cost between $1500 (without light source) to fully integrated systems with a built-in light source at about $3500. Dissecting microscopes costing less than $1500, either of lower quality or with a reduced range of magnifications, may be suitable for some purposes. An adequate 20X or 40X system can be bought for $250-$320.
Scanning and transmission electron microscopes are beyond the reach of all but specialist institutions but are an essential tool for identifying some of the very small algae. None of the algae illustrated here require electron microscopy for identification.
Preserving freshwater algae
Storage and preservation
Algae can be stored initially in a bucket, jar, bottle or plastic bag, with some water from the collecting site. The container should be left open or only half filled with liquid and wide shallow containers are better than narrow deep jars. Note that glass is reportedly not satisfactory for some Chrysophyta and other algae of acidic waters due to its inherent alkalinity damaging cells. However, glass phials are commonly used to collect algae. If refrigerated or kept on ice soon after collecting most algae can be kept alive for short periods (a day or two). If relatively sparse in the sample, some algae can continue to grow in an open dish stored in a cool place with reduced light (traditionally a south-facing window in the Southern Hemisphere).
For long-term storage, specimens can be preserved in liquid (see below), dried, or made into a permanent microscope mount (preferably all three). Even with ideal preservation, examination of fresh material is sometimes essential for an accurate determination. Motile algae particularly must be examined while flagella and other delicate structures remain intact.
Commercial formalin (which is a solution of 40% formaldehyde), diluted between 1/10 and 1/20 with the collecting solution, is the most commonly used fixative. Note that formaldehyde is thought to be carcinogenic and all contact with skin, eyes and air passages should be avoided. FAA (by volume, 40% formaldehyde 1: glacial acetic acid 1: 95% alcohol 8: water 10) or 6-3-1 (by volume, water 6: 90% alcohol 3: 40% formaldehyde 1) solutions give better preservation results for some of the more fragile algae, whereas the standard alcohol and water mix (e.g. 70% ethyl alcohol or industrial methylated spirit) will ruin all but the larger algae.
Algae can be kept in diluted formalin for a number of years, but the solution is usually replaced by 70% ethyl alcohol with 5% glycerin (the latter to prevent accidental drying out).
Lugol's solution is commonly used for short-term (e.g. a few months, but possibly a year or more) storage of microalgae. Dissolve one gram of iodine crystals and two grams of potassium iodide in 300 ml of water. Use three drops of this solution in a 100 ml sample (it should look like very weak tea).
Dried herbarium specimens
Dried herbarium specimens can be prepared by 'floating out' similar to aquatic flowering plants. Ideally, fresh specimens should be fixed prior to drying. Most algae will adhere to absorbent herbarium paper. Smaller, more fragile specimens or tangled, mat-forming algae may be dried onto mica or cellophane. After 'floating out', most freshwater algae should not be pressed but simply left to air dry in a warm dry room. If pressed, they should be covered with a pieces of waxed paper, plastic or muslin cloth so that the specimen does not stick to the drying paper in the press.
To examine a dried herbarium specimen add a few drops of water to the specimen. After a minute or so the specimen will swell and lift slightly from the paper. Carefully remove a small portion of the specimen with forceps or a razer-blade