Clarity

Clarity reflects the amount of dissolved colored or suspended material in any waterbody. Clarity can be affected by natural and introduced materials. Ironically, clarity can be the cause of, as well as a limitation to, productivity. Suspended algae contribute to reduced water clarity, which at the same time can limit light available to growth of algae at depth. In streams and lakes, inorganic sediment can also contribute to reduced clarity. Although usually the case following storms, some waters do maintain high inorganic sediment loads even during baseflow. In the context of nutrient criteria, the utility of clarity measures is related to contributions from suspended algal material in the water column. This material can come from true phytoplankton or from tychoplankton (in rivers, algae dislodged or sloughed from the benthos). In either case, significant correlations have been drawn between the amount of algae in the water column and its clarity, especially for lakes. In fact, one of the common trophic indices for lakes, the Trophic State Index (TSI), can be derived using a measure of transparency (Carlson 1977).

Clarity can be measured in a number of ways. One of the traditional lake quality measures is the vertical Secchi depth transparency measurement, which was adapted from a method developed by an Italian papal advisor, Father Pietro Angelo Secchi, in the 19 th century. Similar horizontal adaptations of transparency measures have been tested for streams and rivers. A similar measure to clarity is turbidity, which measures the scatter and absorption of light by suspended particles. Another way to get at turbidity is to directly measure the total suspended material gravimetrically after filtration. These measures are fairly convenient and, with the exception of gravimetric suspended solids, can be directly measured in the field.

TRANSPARENCY

Transparency is traditionally measured using light meters and is quantified with light extinction coefficients. However, this is rarely done in routine water quality monitoring. For lakes, the most common method is deploying a Secchi disk. There is not common method used for streams and rivers, although a horizontal transparency method has been tested.

Secchi Transparency (Wetzel and Likens 2000)

The Secchi disk is a weighted white or black and white disk, 20cm in diameter that is attached to a graduated line. The disk is lowered over the shaded side of a boat, ideally at midday, and the average of the depths at which the disk disappears and reappears is the estimate of Secchi transparency. Transparency is a function of the reflectance of light from the disk. Any dissolved color or suspended materials that absorb or scatter light will reduce the Secchi transparency, so Secchi depth is proportional to transparency. Transparency measured with Secchi disks is correlated with transmissivity measured directly with photometers, and the Secchi depth is usually around the 10-15% transmission point. Secchi depth in lakes, especially in the absence of color or inorganic suspended material, is highly correlated with algal biomass and can be used to calculate the TSI, which is an accurate measure of lake trophic status (Carlson 1977). Secchi depths range from a few centimeters in very turbid lakes to over 40m in clear, oligotrophic lakes, but most are in the range of 2 to 10m (Wetzel and Likens 2000). There is no comparable routine transparency measure used in streams, however, horizontal black disk samplers have been developed and show promise (Davies-Colley and Smith 2001)

TURBIDITY

Turbidity is a measure of the scatter of light by particles in suspension. It is not the same as clarity and is, in fact, an inverse measure of clarity. Turbidity is caused by suspended particles that intercept light and refract, reflect, or diffract it. These particles include algal cells and high concentrations of water column algae, whether true plankton or dislodged benthic algae (tychoplankton), and will increase turbidity. The Jackson candle turbidimeter was the historical method of choice (units – the Jackson Turbidity Unit or JTU), however nephelometers have replaced those due to their greater sensitivity.

Nephelometry (APHA 1990 Method 2130 B, Wetzel and Likens 2000)

Nephelometers are turbidimeters that measure light scatter at 90º to the incident light beam. Both bench-top and portable nephelometers are available and can give fairly precise and accurate measures, if calibrated properly and regularly. A nephelometer compares the intensity of light scattered by a sample relative to a standard. The greater the scatter, the higher the turbidity, measured as a nephelometric turbidity unit (NTU). Formazin polymer is a commonly used standard reference suspension. Continuous nephelometers, which can be deployed in the field for long periods of continuous turbidity measurement, are available. Turbidities in the range of 0-40 NTU can be measured directly with a nephelometer. Higher values should be diluted to this range, measured, and final concentrations estimated appropriately (APHA 1999).

Literature Cited

American Public Health Association (APHA). 1999. Standard Methods for the Examination of Water and Wastewater, 20 th edition. American Public Health Association, Washington, DC.

Carlson, R.E. 1977. A trophic state index for lakes. Limnology and Oceanography. 22:361-369.

Davies-Colley, R. J. and D. G Smith, 2001. Turbidity, Suspended Sediment, and Water Clarity: A Review. Journal of the American Water Resources Association 37: 1085-1101.

Wetzel, R.G. and G.E. Likens. 2000. Limnological Analyses 3 rd Edition. Springer-Verlag, New York.

Nitrogen is often a limiting nutrient in both marine and freshwater environments, despite preconceived notions about it primarily limiting marine systems (Francoeur et al. 2001). It exists in a number of different valence states, one reason for the variety of biogeochemical reactions that occur as part of the nitrogen cycle. Typical measures of nitrogen include total nitrogen, total organic nitrogen, nitrate, nitrite, and ammonia.

Total nitrogen refers to the total amount of nitrogen in a water sample, which typically contains a variety of inorganic and organic forms. Total nitrogen can be determined by complete oxidation of all forms with a strong oxidant (such as persulfate) and then determination of the nitrate concentration.

Organic nitrogen (including amino acids, peptides, proteins, nucleic acids, and urea) is defined functionally as bound nitrogen in the tri-negative oxidative state, but does not include all organic forms of nitrogen. Kjeldahl nitrogen is a technique that determines organic nitrogen and ammonia (NH3) together. Organic nitrogen can be estimated by subtracting ammonia from total Kjeldahl nitrogen or by subtracting ammonia, nitrate, and nitrite from a measure of total nitrogen. Organic nitrogen occurs in the range of tens of ug N/L to more than 20 mg N/L in raw sewage.

Total oxidized nitrogen is the sum of nitrate and nitrite. Nitrate () is the most oxidized form of nitrogen and is usually found in low concentrations in oligotrophic waters (10-100 ug -N /L), but can be in the 10-100 mg -N/L range in biologically treated effluent or eutrophic waters. Nitrate is taken up by plants and algae and enzymatically reduced to the organic amino form with nitrate reductase. Nitrate can also be reduced by denitrification, a microbially-mediated process occurring under anaerobic conditions that uses nitrate as a terminal electron acceptor and resulting in the production, ultimately, of nitrogen gas (N2). Nitrite () is an intermediate oxidative state that is usually found in low concentrations in water. However, it is used in some industrial applications, the effluent of which can contain high concentrations.

Ammonia (NH3) is the most reduced form of nitrogen. It originates naturally from the decomposition of organic nitrogen compounds and hydrolysis of urea. It is usually present in low concentrations tens of ug N/L, except where enrichment from various sources results in concentrations in the hundreds of ug NH3-N/L to tens of mg NH3-N/L range.

Concentrations of nitrogen ions are usually reported as elemental nitrogen in its various forms and -N, -N, and NH3-N are technically “N as nitrate, N as nitrite, and N as ammonia”. Care should be made to check that concentrations are given in terms of the elemental nitrogen concentration.

The following are brief descriptions of some standard methods for measuring different forms of nitrogen. Users interested in further specific and detailed methods are asked to refer to the literature cited.

TOTAL NITROGEN

Total nitrogen methods employ strong oxidants to convert all forms of nitrogen (reduced and oxidized, bound and dissolved) into nitrate ions. It differs from total Kjeldahl nitrogen, which does not measure oxidized forms of nitrogen.

Persulfate Digestion (APHA 1999 Method 4500-N C.)

The persulfate method, as mentioned above, uses the strong oxidant, persulfate, to convert all forms of nitrogen in a sample into the nitrate molecule, which most efficiently occurs at 100 to 110 degrees C in an alkaline environment. This is usually done with digestion tubes in autoclaves, hotplates, etc. The resulting nitrate is then measured using nitrate methods (see below) after cooling and buffering the sample.

ORGANIC NITROGEN

Organic nitrogen methods measure principally nitrogen in the tri-negative state (amino). They do not measure other organic forms of nitrogen (e.g., -azide, -azine, -azo, nitro, etc.). Organic nitrogen concentration can be elevated in areas with large potential sources of organic nitrogen without treatment and in areas with high levels of nitrogen inputs in general. Even in oligotrophic areas, organic forms may predominate the nitrogen pool. The traditional method for measuring organic nitrogen is the Kjeldahl method.

Kjeldahl Method (APHA 1999 Method 4500-N org B)

This method measures organic-N and ammonia. The principle of this method is that amino-nitrogen compounds are converted to ammonium in the presence of sulfuric acid, potassium sulfate, and cupric sulfate during a digestion. Free ammonia (NH3) is also converted into ammonium (). After the initial digestion, base is added and the sample distilled to remove the ammonium. Ammonium is then measured using an appropriate ammonium method (e.g., phenate method). As this method commonly measures dissolved ammonia as well as organic-N, organic-N alone can be measured by removing the ammonia first and using a pre-distillation or subtracting the ammonia measured using a dissolved ammonia method (see below). This method is applicable over a wide range of organic-N plus ammonia concentrations but requires large volumes for low concentration waters. The macro-Kjeldahl method typically uses 800 ml Kjeldahl flasks. A micro-Kjeldahl method exists that is applicable over the range of organic plus ammonia nitrogen of 0.2 to 2 mg N/L (APHA 1999 4500-N org C). This method simply uses smaller volume (100 ml) Kjeldahl flasks.

AMMONIA NITROGEN

Ammonia is the most reduced form of nitrogen. It usually exists in low concentrations and most often exists as the ammonium ion () except under high pH (>9.0)(Wetzel and Likens 2000). Ammonia can generally be measured directly in surface water samples, although filtration can be used as well as distillation for samples with common interference or very high concentrations (> 5 mg NH3-N/L)(APHA 1999).

Phenate Method (APHA 1999 Method 4500-NH3 C, Wetzel and Likens 2000)

The principal of the phenate method is that ammonium, in the presence of hypochlorite and phenol forms an intense blue compound, indophenol, that can be measured spectrophotometrically. The amount of indophenol produced is proportional to the ammonium concentration. It is a fairly straightforward method, which is accurate to very low concentrations (0.01 mg NH3-N/L) with long path cell lengths and is linear up to 0.6 mg NH3-N/L. Samples above this concentration may have to be diluted. There is an automated form of the phenate method (APHA 1999 Method 4500-NH3 G), which follows the same principal as the manual method, but uses continuous flow analytical machines that automate the process and can be used for analyzing samples in large batches. The automated method is applicable over the range of 0.02 to 2.0 mg NH 3-N/L.

NITRITE NITROGEN

Nitrite is the trivalent form of nitrogen which usually exists in concentrations below that of nitrate. However, nitrite concentrations can be elevated, for example, in rivers under warm, slow-moving conditions or in anaerobic lake strata as a result of high rates of denitrification (dissimilatory nitrate reduction or denitrification)(Wetzel 2001), or in effluent from industrial applications employing nitrite.

Colorimetric Method (APHA 1999 Method 4500- A, Wetzel and Likens 2000)

The principal of this method is that nitrite forms a reddish purple azo dye under acidic conditions when combined with certain reagents. This color is produced in proportion to the concentration of nitrite and can be measured spectrophotometrically. This is the common nitrite method and is useful in the range of 10 to 1000 mg -N/L. Lower concentrations can be estimated by using a 5-cm path cell. Higher concentrations should be diluted.

NITRATE NITROGEN

Nitrate is the most oxidized common form of nitrogen in freshwaters. It is taken up by algae and plants and reduced to the amino form (assimilatory nitrate reduction) and by denitrifying bacteria and converted into nitrogen gas (dissimilatory nitrate reduction or denitrification). Nitrate concentration can be quite high where nitrogen loading exists as a result of direct nitrate input or as a result of nitrification of reduced organic forms by bacteria. Nitrate measurement is difficult because of the relatively complex method and apparatus required, common interferences, and the limited range of different methods. Nitrate can be determined by ion chromatography or capillary ion electrophoresis, which are fairly straightforward. The more traditional technique is the cadmium reduction method.

Cadmium Reduction Method (APHA 1999 Method 4500- E, Wetzel and Likens 2000)

The principal of this method is that nitrate is reduced almost completely to nitrite in the presence of cadmium. The nitrite produced is then determined following the standard colorimetric method described for nitrite above. Note that this method measures both nitrate and nitrite in the sample. Nitrite can be subtracted by measuring a subsample directly without the reduction step. The manual method is applicable across nitrate ranges from 0.01 to 1 mg -N/L. Higher concentrations can be diluted. An automated version of this method exists (APHA 1999 Method 4500- F), which follows the same principal as the manual method, but uses continuous flow analytical machines that automate the process and can be used for analyzing samples in large batches. The automated method is applicable over the range of 0.001 to 10.0 mg -N /L and higher concentrations can be diluted.

Literature Cited

American Public Health Association (APHA). 1999. Standard Methods for the Examination of Water and Wastewater, 20 th edition. American Public Health Association, Washington, DC.

Francoeur, S.N. 2001. Meta-analysis of lotic nutrient amendment experiments: detecting and quantifying subtle responses. Journal of the North American Benthological Society 20:358-368.

Wetzel, R.G. 2001. Limnology: Lake and River Ecosystems 3 rd edition. Academic Press, New York.

Wetzel, R.G. and G.E. Likens. 2000. Limnological Analyses 3 rd Edition. Springer-Verlag, New York.

Periphyton (or aufwuchs) is the community of organisms including plants, bacteria, fungi, protozoa, and invertebrates living attached to submerged substrates (Dodds 2002). These communities compose the biofilms in waterbodies where benthic algae reside. As a result, they are the focus of sampling for algal response to nutrients and are, therefore, important in nutrient criteria development. A variety of substrates support periphyton communities and are named accordingly: epilithon (on rocks), epiphyton (on plants), epidendron or epixylon (on wood), epipsammon (on sand), epipelon (on fine sediments), and epizoon (on aquatic animals).

Two basic attributes of periphyton are most often sampled: biomass and composition. Biomass is generally measured by removing periphyton from substrates and weighing the resultant organic matter, extracting the chlorophyll to provide a relative estimate of algal abundance, or counting the number of algal cells and using a biovolume conversion to estimate biomass. Algal assemblage composition typically consists of removing periphyton, preserving it, and then identifying the taxa to the lowest possible taxonomic level. However, rapid periphyton methods have been developed for doing field based community assessment. Both biomass and composition can be measured from the same field samples, which are split into a subsample for biomass and one for community composition.

Below is a brief review of common methods. Interested readers are encouraged to go to the original literature for more detailed methodologies and greater background information.

SAMPLE COLLECTION

Periphyton samples can be collected using quantitative or semi-quantitative methods and a variety of programs have developed specific methods (e.g., Barbour et al. 1999, Moulton et al. 2002). Quantitative methods consist of sampling a known area of substrate, usually with a sampling device that can isolate an area which can be removed (e.g., a petri dish/spatula on sand) in situ or by removing substrates and removing the material from a specific area. Either specific substrates can be targeted or multi-habitat samples can be taken and composited. Known area samples are best for biomass estimates; but semi-quantitative, multi-habitat samples may gather more taxa. Passive samplers can also be deployed for some period of colonization. Glass slides, the top plate of Hester-Dendy macroinvertebrate samplers, and clay tiles are all examples of samplers that have been used as passive sampling substrates for periphyton. The advantage of these is the standardized sampling area and substrate characteristics. The major disadvantage is the fact that these artificial substrates may not accurately reflect the true periphyton community.

ALGAL BIOMASS

Algal biomass refers to the mass of algal material within the periphyton. It can be measured gravimetrically (weighed), using chlorophyll extractions, or by converting cell count data to mass using biovolume conversions. Gravimetric measures rely on weighing the amount or organic matter within the periphyton. As it is generally prohibitive to separate algal and non-algal material for routine analysis, the mass [usually the mass after combustion called ash-free dry mass (AFDM)] can include substantial non-algal material. Some people use AFDM to chlorophyll ratios to look at the proportion of non-algal material, but this is only an approximation. The content of chlorophyll per cell varies by a number of factors: species, nutrient conditions, light condition, and temperature. Therefore, chlorophyll is not a direct measure of biomass, however it has long been used as a relative measure of algal abundance (Wetzel 2001). Cell biovolumes vary by taxa, their shapes, and a number of other factors affecting cellular composition. However, if quantitative genus or species level data are collected, then biomass can generally be estimated by applying shape specific geometric volume conversions.

Ash-free dry mass (Hauer and Lamberti 1996, APHA 1999 Method 10300 C)

After scraping a known area, the resulting “scrapate” is diluted to a known volume and a subsample of that volume is either weighed in pre-combusted and pre-weighed crucible or weighed following filtration through appropriate pre-weighed and pre-combusted glass fiber filters (0.5 to 0.7 um pore size). Crucibles and filters are then dried, weighed for dry mass, and then combusted to remove all organic matter (e.g., 500º C) and re-weighed to estimate ash-free dry mass. Samples for AFDM can be field filtered or brought to a lab for processing. Sample preservation for AFDM is not straightforward, but samples should be handled to limit respiratory loss of organic matter (e.g., placed on ice and frozen (-20 to -60° C) if there will be a delay in processing). Consult the different specific method for more on sample preservation. Values are expressed as g AFDM per unit area.

Pigment analysis (Hauer and Lamberti 1996, APHA Method 1990 10200 H, 10300 C)

A sub-sample of material removed from substrates and usually filtered onto glass fiber filters (0.5 to 0.7 µm pore size) is used for pigment analysis as an indicator of algal biomass in periphyton. The composition of photosynthetic pigments varies by algal taxa, but chlorophyll a is the principal pigment used because it is the most common and abundant. Chlorophylls b and c and the carotenoids absorb light at different wavelengths and can be quantified as well, if desired. Chlorophyll a degrades into different phaeopigments that absorb at the same wavelength as chlorophyll a. For this reason, chlorophyll concentration is usually estimated in a sample after extraction and then acidified to convert all the chlorophyll to phaeopigments and chlorophyll a determined by difference. Chlorophyll a is historically and most commonly measured using spectrophotometry, although fluorometry is more sensitive and becoming more commonly used.

Chlorophyll and other photopigments must be extracted from algae to quantify biomass using pigment analysis. It is best to work in low light to avoid degradation. While a variety of extraction solvents exist, alkaline aqueous acetone is still the standard recommended method (APHA 1999). Algae retained on glass fiber filters (0.5 to 0.7 µm pore size) are typically ground in aqueous acetone, allowed to sit for a period of time for extraction in the dark and under refrigeration, and the ground filter solution is centrifuged to separate the filter from the pigments. Pigments are then decanted and measured directly either with spectrophotometry or fluorometry. The flourometric method is usually standardized using spectrophotometric measurement of a known chlorophyll concentration. Fluorometry is more sensitive to low concentrations, and should give comparable estimates. Consult an appropriate methods source for details on the use of either of these detection methods, including appropriate detectors or wavelengths.

Preservation of samples for chlorophyll is important. Filtration should occur as soon as possible (Wetzel and Likens 2000) and chlorophyll should be extracted from filters as soon as possible. Field filtration is ideal. Addition of magnesium carbonate either to the sample solution or on top of the filter after filtration has been advocated to reduce acidity, since acidity degrades chlorophyll. Filters that will not be analyzed immediately, should be folded in half, placed into labeled dark containers (i.e., wrapped in aluminum foil), and placed on ice or immediately frozen. Pigment samples may be frozen for a few days (at -20 to -60° C), but should be analyzed as soon as possible, since degradation of chlorophyll does occur (Wetzel and Likens 2000).

Biovolume (Wetzel and Likens 2000, APHA 1999 10200 I)

Biovolume is determined by multiplying the number of each taxon identified in a sample by the average biovolume of that taxon, calculated using equations of geometric shapes most approximating that taxon, and summing these values across all the taxa in a sample. Equations for many different taxa have already been developed (Hillebrand et al. 1999) or general equations may be applied (Wetzel and Likens 2000). This approach obviously requires more effort, since individual taxa must not only be counted, but average dimensions calculated for each as well (average of 20 individuals is recommended, APHA 1999). This method does, however, offer one of the more accurate measures of live algal biomass.

Visual Estimate (Stevenson and Bahls 1999)

As part of a field-based rapid periphyton survey developed for use in the Rapid Bioassessment Protocols, a quick visual estimation of algal biomass was developed. This approach uses gridded view buckets to visually estimate macroalgal biomass and microalgal cover. While not as accurate as actual measures of algal biomass, the technique does allow rapid relative estimates of composition and standing crop or biomass.

ALGAL COMPOSITION

The composition of the algal assemblage in stream periphyton can provide information about the physical and chemical environment. Unlike simple grab samples of water, the assemblage of algae integrate water quality conditions over long periods of time. Each taxon has specific optima for the wide variety of physical and chemical conditions to which periphyton are exposed, including sensitivities or tolerance to nutrients, acidity, temperature, oxygen, silt, and other variables. The average environmental conditions select which taxa survive and thrive in a stream. As a result, a great deal of environmental information can be inferred from which algae are found in the periphyton, including nutrient conditions. The composition of algae is usually determined with microscopy, and most taxa can be identified to species by well trained taxonomists. A rapid visual estimate of rough benthic algal composition has also been developed.

Microscopy (Wetzel and Likens 2000, APHA 1999 10300C, Stevenson and Bahls 1999)

Unfiltered periphyton samples collected from the stream need to be preserved. The preferred preservative is Lugol’s solution, but other optional preservatives include 2% M3 fixative, 4% buffered formalin, or 2% glutaraldehyde. The sample is typically homogenized with a tissue homogenizer and placed into a specific cell for viewing (e.g., Palmer-Maloney or Sedgwick-Rafter cells) of large algal taxa. These can be enumerated a number of ways, but it can be difficult to enumerate colonial or filamentous individuals (Stevenson and Bahls 1999, Wetzel and Likens 2000), and magnification is limited for these sample cells. Sedimentation chambers can also be used for identifying smaller cells if inverted microscopes are available. In this approach, a known subsample of the homogenized sample is placed into a sedimentation chamber, where the algae settle. The chamber is then placed on an inverted microscope, where greater magnification can be used to identify taxa. For diatom analysis, it is necessary to remove organic matter that might otherwise interfere with identification. This can be done with combustion or chemical oxidation of subsamples, followed by slide mounting with a suitable highly refractive medium to make permanent slides. Compound microscopy at 1000x under oil immersion is then used to identify diatoms to species. From these data, accurate identifications of the resident algal periphyton assemblage can be made. When combined with autecological information for the different algal taxa, a number of inferences about the integrated physical and chemical conditions of a stream or lake can be made.

Rapid visual estimates (Stevenson and Bahls 1999)

The rapid visual approach uses gridded view buckets to characterize periphyton assemblages. Very coarse-level taxonomic information can be collected by individuals trained to recognize specific macro-algal taxa. The buckets are placed randomly along transects and the algal assemblage identified to specific macroalgal genera and/or to coarse algal groups (diatoms or blue-green algae).

Literature Cited

American Public Health Association (APHA). 1999. Standard Methods for the Examination of Water and Wastewater, 20 th edition. American Public Health Association, Washington, DC.

Barbour, M.T., J. Gerritsen, B.D. Snyder, and J.B. Stribling. 1999. Rapid Bioassessment
Protocols for Use in Streams and Wadeable Rivers: Periphyton, Benthic
Macroinvertebrates and Fish, Second Edition. EPA 841-B99-002. US Environmental
Protection Agency, Office of Water, Washington, D.C.

Dodds, W.K. 2002. Freshwater Ecology: Concepts and Environmental Applications. Academic Press, New York.

Hauer, F.R. and G.A. Lamberti. 1996. Methods in Stream Ecology. Academic Press, New York.

Hillebrand, H., C.-D. Durselen, D. Kirschtel, U. Pollingher, and T. Zohary. 1999. Biovolume calculation for pelagic and benthic microalgae. Journal of Phycology 35: 403-424.

Moulton II, S.R., J.G. Kennen, R.M. Goldstein, and J.A. Hambrook, 2002. Revised Protocols for Sampling Algal, Invertebrate, and Fish Communities as Part of the National Water-Quality Assessment Program. U.S. Geological Survey Open-File Report 02-150.

Stevenson, R.J. and L.L. Bahls. 1999. Periphyton protocols. In Barbour, M.T., J. Gerritsen, B.D. Snyder, and J.B. Stribling. 1999. Rapid Bioassessment
Protocols for Use in Streams and Wadeable Rivers: Periphyton, Benthic
Macroinvertebrates and Fish, Second Edition. EPA 841-B99-002. US Environmental
Protection Agency, Office of Water, Washington, D.C.

Wetzel, R.G. 2001. Limnology: Lake and River Ecosystems 3 rd edition. Academic Press, New York.

Wetzel, R.G. and G.E. Likens. 2000. Limnological Analyses 3 rd Edition. Springer-Verlag, New York.

Phosphorus is also a common limiting nutrient in both marine and freshwater systems. Unlike nitrogen, phosphorus exists predominantly as the phosphate ion in freshwater – . Phosphates are classified as orthophosphates (single phosphate ions), condensed phosphates (polyphosphates), and organically bound phosphates. Phosphates occur as dissolved or bound forms – either in organic particles or attached to detritus or inorganic sediment. Phosphine gas (PH 3) is rare. Phosphorus in surface waters comes from a variety of sources. Natural sources include bedrock and precipitation. Common anthopogenic sources include fertilizers and detergents, as well as human waste and food residues (APHA 1999).

Phosphorus is divided into particulate and “dissolved” forms, the latter of which are defined principally based on the method selected. Figure 1 illustrates how the different fractions are measured. The total phosphorus fractions consist of particular and “dissolved” fractions, measured from an unfiltered sample. Total fractions include phosphate bound in organic and to inorganic particles. Dissolved fractions are those measured after filtration through a membrane filter. Dissolved organic forms include phospho-lipids, nucleic acids, and ATP.

Phosphate analysis consists principally of converting the various forms of phosphorus to dissolved orthophosphate () followed by direct colorimetric measurement. The separation of the different forms (Figure 1) is a-nalytically defined but these have been selected to be consistent with more or less functional P fractions (APHA 1999). The separation of dissolved and particulate fractions has traditionally been through 0.45 mm membrane filters. This distinction is not absolute, but simply allows for a relatively precise and replicable standard of measurement (APHA 1999). Within the dissolved and total phosphorus fractions, phosphorus is further divided into three principal forms of phosphorus: reactive, acid-hydrolyzable, and organic phosphorus.

Reactive phosphorus, in both dissolved and particulate fractions, is that P which reacts with reagents in colorimetric tests without hydrolysis or digestion (APHA 1999). While reactive phosphorus is principally orthophosphate, it also includes a small fraction of condensed phosphate which is hydrolyzed during the analysis.

Acid hydrolysis is used to convert all dissolved and particulate condensed forms into orthophosphate. Again, this process likely liberates some organically bound phosphorus and may include some of that fraction, but this can be minimized procedurally (APHA 1999). This analytical artifact is the basis for calling the fraction “acid-hydrolyzable” rather than “condensed phosphate”.

Lastly, organic and organically-bound phosphorus is that fraction released only by oxidative digestion. Organic P also occurs in dissolved and particulate forms.

Suspended fractions are usually determined by difference (Figure 1).

Sample Storage

Filtration for dissolved analysis, if desired, usually occurs during or immediately after sample collection. Samples can be frozen or chemically preserved. Equipment type and lab environment can affect P analysis, so extra care must be taken. Preparation of equipment used in P collection and analysis is strict and must be adhered to minimize contamination and sample error.

ACID HYDROLYSIS AND DIGESTION

Digestions are used to estimate the total phosphorus and total organic phosphorus fractions (by subtraction). There are three principal digestion techniques: perchloric, nitric acid-sulfuric acid, and persulfate. Perchloric acid digestion is recommended only for the most difficult samples (e.g., sediments) and is time-consuming and severe (APHA 1999). Nitric acid-sulfuric acid digestion is appropriate for most samples. Persulfate digestion is the simplest technique and should be checked against the other two for comparability (APHA 1999).

Acid Hydrolysis (APHA 1999 Method 4500-P B2)

Acid hydrolysis is used for measuring the acid-hydrolyzable fraction and is defined as the difference between the concentrations in an untreated sample (reactive P) and one treated with mild acid. It includes condensed phosphates and potentially some organic phosphate compounds (APHA 1999). This method involves acidifying a sample with sulfuric and nitric acids followed with gentle boiling. The orthophosphate liberated is then measured using one of the colorimetric methods (see below).

Perchloric Acid Digestion (APHA 1999 Method 4500-P B3)

This fairly intense procedure involves acidifying a sample with nitric acid and then digesting the sample in a solution of nitric acid and perchloric acid. This is then neutralized with sodium hydroxide. Orthophosphate is then measured using one of the colorimetric methods (see below).

Sulfuric Acid-Nitric Acid Digestion (APHA 1999 Method 4500-P B4)

This method uses a digestion rack much like those used for micro-Kjeldahl nitrogen determination. In this approach, sulfuric and nitric acids are added to a sample and digested. The orthophosphate liberated is then measured using one of the colorimetric methods (see below).

Persulfate Digestion (APHA 1999 Method 4500-P B5, Wetzel and Likens 2000)

In this approach, persulfate is added to a pH-adjusted sample and boiled for a set period of time. The orthophosphate liberated is then measured using one of the colorimetric methods (see below).

COLORIMETRIC METHODS FOR PHOSPHATE

The colorimetric methods are fairly similar and depend principally on the range of concentrations desired. The vanadomolybdophosphoric acid method is used for phosphate concentrations between 1 and 20 mg P/L and the stannous chloride and ascorbic acid methods are used for ranges between 0.01 and 6 mg P/L. Extractions and longer cell paths may improve detection on the lower ranges. Ion chromatography and capillary ion electrophoresis methods can also be used for determining orthophosphate concentrations in undigested samples.

Vanadomolybdophosphoric Acid Method (APHA 1999 Method 4500-P C)

The principle of this test is that ammonium molybdate reacts with orthophosphate under acidic conditions and in the presence of vanadium to form a yellow color which is proportional to the concentration of phosphate in the sample. This color can then be measured with a colorimeter and the phosphate concentration of the sample estimated. Again, this method is generally recommended over the 1 to 20 mg P/L range.

Stannous Chloride Method (APHA 1999 Method 4500-P D)

In this method, molybdophosphoric acid (formed by the reaction of orthophosphate with ammonium molybdate under acidic conditions) is reduced by stannous chloride to form a blue molybdenum color which is proportional to the concentration of phosphate in the sample. This color can then be measured with a colorimeter and the phosphate concentration of the sample estimated. With long path cells, this method can measure P down to 0.007 mg P/L. An extraction step using separation with benzene-isobutanol can increase the sensitivity for low concentration samples.

Ascorbic Acid Method (APHA 1999 Method 4500-P E, Wetzel and Likens 2000)

In this method, ammonium molybdate and potassium antimonyl tartrate react with orthophosphate under acidic conditions to form phosphomolybdic acid which is reduced to a blue color solution by ascorbic acid. The blue color formed is proportional to the concentration of phosphate in the sample. This color can then be measured with a colorimeter and the phosphate concentration of the sample estimated. The method is accurate to 0.01 mg P/L with a 5 cm cell path length. An extraction step using separation with solvent can increase the sensitivity for low concentration samples. An automated version of this method exists (APHA 1999 Method 4500- P F), which follows the same principal as the manual method, but uses continuous flow analytical machines that automate the process and can be used for analyzing samples in large batches. The automated method is applicable over the range of 0.001 to 10.0 mg P /L. Higher concentrations can be diluted.

Literature Cited

American Public Health Association (APHA). 1999. Standard Methods for the Examination of Water and Wastewater, 20 th edition. American Public Health Association, Washington, DC.

Francoeur, S.N. 2001. Meta-analysis of lotic nutrient amendment experiments: detecting and quantifying subtle responses. Journal of the North American Benthological Society 20:358-368.

Wetzel, R.G. 2001. Limnology: Lake and River Ecosystems 3 rd edition. Academic Press, New York.

Wetzel, R.G. and G.E. Likens. 2000. Limnological Analyses 3 rd Edition. Springer-Verlag, New York.

Phytoplankton

Phytoplankton is the assemblage of autotrophs found in the water column including diatoms, green algae, cyanobacteria, dinoflagellates, and other algal taxa.  The phytoplankton includes single celled as well as colonial and filamentous forms; and it includes both true phytoplankton as well as periphytic algae that have been dislodged from the benthos (tychoplankton), especially in rivers.  In waterbodies with sufficient residence time where true phytoplankton can develop (e.g., lakes, estuaries, and large rivers), this assemblage is the focus of sampling for algal response to nutrients and is, therefore, important in nutrient criteria development.  There are well established relationships between nutrient loading and phytoplankton biomass in lakes (Wetzel 2001).  In rivers, the same relationships have been explored only less so and the relationships are often far more variable than those for lakes.

As with periphyton, two basic attributes of phytoplankton are most often sampled: biomass and composition.  Biomass is generally measured by filtering phytoplankton and weighing the resultant organic matter, extracting the chlorophyll to provide a relative estimate of algal abundance, or counting the number of algal cells and using a biovolume conversion to estimate biomass.  Phytoplankton assemblage composition typically consists of preserving a representative sample of water and identifying the algal taxa to the lowest possible taxonomic level.  Both biomass and composition can be measured from the same field samples, which are split into a subsample for biomass and one for assemblage composition.

The following is a brief review of common methods.  Interested readers are encouraged to examine the original literature for more detailed methodologies and greater background information.

SAMPLE COLLECTION

Phytoplankton samples can be collected using quantitative or semi-quantitative methods.  Quantitative methods consist of sampling a known volume of water, usually with a sampling device that can isolate a known volume at a specific depth (e.g., Van Dorn, Niskin, Nansen, or Kemmerer Bottle).  Another option is to use a long, straight tubular sampler which allows the collection of a depth integrated sample, essentially a “core” of the water column which collects plankton over a range of depths.

ALGAL BIOMASS

Algal biomass refers to the mass of algal material within the phytoplankton.  It can be measured gravimetrically (weighed), using chlorophyll extractions, or by converting cell count data to mass using biovolume conversions.  Gravimetric measures rely on weighing the amount or organic matter within the phytoplankton.  As it is generally prohibitive to separate algal and non-algal material for routine analysis, the mass [usually the mass after combustion called ash-free dry mass (AFDM)] can include substantial non-algal material, although this is generally more severe in lotic than lentic systems.  Some people use AFDM to chlorophyll ratios to look at the proportion of non-algal material, but this is only an approximation.  The content of chlorophyll per cell varies by a number of factors: including species, nutrient conditions, light condition, and temperature.  Therefore, chlorophyll is not a direct measure of biomass, however it has long been used as a relative measure of algal abundance (Wetzel 2001).  Cell biovolumes vary by taxa, their shapes, and a number of other factors affecting cellular composition.  However, if quantitative genus or species level data are collected, then biovolume can generally be estimated by applying shape specific geometric volume conversions.

Ash-free dry mass (Wetzel and Likens 2000, APHA 1999 Method 10200 I)

A subsample of the original phytoplankton sample is either weighed in pre-combusted and pre-weighed crucibles or weighed following filtration through appropriate pre-weighed and pre-combusted glass fiber filters (0.5 to 0.7 µm pore size).  Crucibles and filters are then dried, weighed for dry mass, and then combusted to remove all organic matter (e.g., 500º C) and re-weighed to estimate AFDM.  Samples for AFDM can be field filtered or brought to the lab for processing there.  Sample preservation for AFDM is not straightforward, but samples should be handled to limit respiratory loss of organic matter (e.g., placed on ice and frozen (-20 to -60° C) if there will be a delay in processing).  Consult the different specific method for more on sample preservation.  Values are expressed as g AFDM per unit volume.

Pigment analysis (Wetzel and Likens 2000, APHA 1990 Method 10200 H)

A subsample of the phytoplankton sample is usually filtered onto glass fiber filters (0.5 to 0.7 µm pore size) and used for pigment analysis as an indicator of algal biomass.  On average, chlorophyll constitutes 1.5% of the AFDM of algae, but the ratio of chlorophyll to cell biomass varies by taxa, so biomass estimates using this technique should be considered relative.  The composition of photosynthetic pigments also varies by algal taxa, but chlorophyll a is the principal pigment used because it is the most common and abundant.   Chlorophylls b and c and the carotenoids absorb light at different wavelengths and can be quantified as well, if desired.

Chlorophyll a degrades into different phaeopigments that absorb at the same wavelength as chlorophyll a.  For this reason, chlorophyll concentration is usually estimated in a sample after extraction and then acidified to convert all the chlorophyll to phaeopigments and chlorophyll a determined by difference. Chlorophyll a is historically and most commonly measured using spectrophotometry, although fluorometry is more sensitive and becoming more commonly used. 

Chlorophyll and other photopigments must be extracted from algae to quantify biomass using pigment analysis.  It is best to work in low light to avoid degradation.  While a variety of extraction solvents exist, alkaline aqueous acetone is still the standard recommended method (APHA 1999).  Algae retained on glass fiber filters (0.5 to 0.7 um pore size) are typically ground in aqueous acetone, allowed to extract for a period of time under refrigeration in the dark, and the ground filter solution is centrifuged to separate the filter from the pigments.  Pigments are then decanted and measured directly either with spectrophotometry or fluorometry.  The flourometric method is usually standardized using spectrophotometric measurement of a known chlorophyll concentration and while fluorometry is more sensitive, should give comparable estimates. Consult an appropriate methods source for details on the use of either of these detection methods, including appropriate detectors, wavelengths, etc.

Preservation of samples for chlorophyll is important.  Filtration should occur as soon as possible (Wetzel and Likens 2000) and chlorophyll should be extracted from filters as soon as possible. Field filtration is ideal. Addition of magnesium carbonate either to the sample solution or on top of the filter after filtration has been advocated to reduce acidity, since acidity degrades chlorophyll.  Filters that will not be analyzed immediately, should be folded in half, placed into labeled dark containers (e.g., wrapped in aluminum foil), and placed on ice or immediately frozen.  Pigment samples may be frozen for a few days (at -20 to -60 degree C), but should be analyzed as soon as possible, since degradation of chlorophyll does occur (Wetzel and Likens 2000).

Biovolume (Wetzel and Likens 2000, APHA 1999 Method 10200 I)

Biovolume is determined by multiplying the number of each taxon identified in a sample by the average biovolume of that taxon, calculated using equations of geometric shapes most approximating the shape of each taxon, and summing these values across all the taxa in a sample.  Equations for many different taxa have already been developed (Hillebrand et al. 1999) or general equations may be applied (Wetzel and Likens 2000).  This approach obviously requires more effort, since individual taxa must not only be counted, but average dimensions calculated for each as well (average of 20 individuals is recommended, APHA 1999).  This method does, however, offer one of the more accurate measures of live algal biomass. 

ALGAL COMPOSITION

The composition of the algal assemblage of phytoplankton can provide information about the physical and chemical environment.  Unlike simple grab samples of water, the assemblage of algae integrate water quality conditions over long periods of time.  Each taxon has specific optima for the wide variety of physical and chemical conditions to which periphyton are exposed, including sensitivities or tolerance to nutrients, acidity, temperature, oxygen, silt, etc.  The average environmental conditions select which taxa survive and thrive.   As a result, a great deal of environmental information, including nutrient conditions, can be inferred from which algae are found in the phytoplankton.  This approach was developed by paleolimnologists, who identify the diatoms found in sedimentary strata of lakes to infer historical, even ancient lake environmental conditions based on this principle.  The composition of algae is usually determined with microscopy, and most taxa can be identified to species by well trained taxonomists. 

Microscopy (Wetzel and Likens 2000, APHA 1999 Method 10200)

Unfiltered phytplankton samples need to be preserved.  The preferred preservative is Lugol’s solution, but other optional preservatives include 2% M3 fixative, 4% buffered formalin, or 2% glutaraldehyde.  Phytoplankton samples containing macroalgae can be homogenized with a tissue homogenizer and placed into a specific cell (e.g., Sedgwick-Rafter (up to 200x) or Palmer-Maloney (up to 500x) cells) for viewing larger algal taxa.  These can be enumerated a number of ways, but it can be difficult to enumerate colonial or filamentous individuals (APHA 1999, Wetzel and Likens 2000).  Magnification is limited with these cells, so microphytoplankton often cannot be identified accurately.  Sedimentation chambers and inverted microscopes (500-600x) are ideal for identifying large and smaller cells together.  In this approach, a known subsample of the homogenized sample is placed into a sedimentation chamber, where the algae settle.  The chamber is then placed on an inverted microscope, where higher objectives may be used.  Ocular grids may improve counting efficiency.  For the highest magnification (1000x), upright microscopes with oil immersion lenses are required.  These are typically used for small diatom analysis.  For diatom analysis, it is necessary to remove organic matter that might otherwise interfere with identifying diatoms.  This can be done with combustion or chemical oxidation of subsamples, followed by slide mounting with a suitable highly refractive medium to make permanent slides.  Other options include membrane filtration followed by clearing of the membranes.  Compound microscopy at 1000x under oil immersion is then used to identify diatoms to species.  Again, use of ocular grids may increase counting efficiency considerably.  From these samples, accurate identification of the resident phytoplankton assemblage can be made.  When combined with autecological information for the different algal taxa, a number of inferences about the integrated physical and chemical conditions of a lake can be made.

Literature Cited

American Public Health Association (APHA).  1999.  Standard Methods for the Examination of Water and Wastewater, 20th edition.  American Public Health Association, Washington, DC.

Hillebrand, H., C.-D. Durselen, D. Kirschtel, U. Pollingher, and T. Zohary.  1999.  Biovolume calculation for pelagic and benthic microalgae.  Journal of Phycology 35: 403-424.

Wetzel, R.G.  2001.  Limnology: Lake and River Ecosystems 3rd edition.  Academic Press, New York.

Wetzel, R.G. and G.E. Likens.  2000.  Limnological Analyses 3rd Edition.  Springer-Verlag, New York.