Biological test method for measuring the inhibition of growth using freshwater macrophyte: appendices

Appendices

1. Members of the Inter-Governmental Environmental Toxicity Group (as of December 2006)
2. Environment Canada, Environmental Protection Service, Regional and Headquarters Offices
3. Procedural Variations for Culturing Lemna spp., and for Undertaking Growth Inhibition Tests Using Lemna spp., as Described in Canadian, American, and European Methodology Documents
4. Review of Culture and Test Media Used in Lemna spp. Growth Inhibition Tests, as Described in Canadian, American, and European Methodology Documents
5. General Description of Lemna minor
6. Axenic Culture Techniques for Lemna (Acreman, 2006)
7. Logarithmic Series of Concentrations Suitable for Toxicity Tests
8. Biological Test Methods and Supporting Guidance Documents Published by Environment Canada’s Method Development & Applications Section

Appendix A: Members of the Inter-Governmental Environmental Toxicity Group (as of December 2006)

Federal, Environment Canada

W. Antoniolli
Environmental Protection Service
Edmonton, Alberta

C. Blaise
Centre St. Laurent
Montreal, Quebec

U. Borgmann
National Water Research Institute
Burlington, Ontario

J. Bruno
Pacific Environmental Science Centre
North Vancouver, British Columbia

C. Buday
Pacific Environmental Science Centre
North Vancouver, British Columbia

K. Doe
Atlantic Environmental Science Centre
Moncton, New Brunswick

G. Elliott
Environmental Protection Service
Edmonton, Alberta

F. Gagné
Centre St. Laurent
Montreal, Quebec

M. Harwood
Environmental Protection Service
Montreal, Quebec

S. Hendry
Environmental Technology Centre
Ottawa, Ontario

D. Hughes
Atlantic Environmental Science Centre
Moncton, New Brunswick

P. Jackman
Atlantic Environmental Science Centre
Moncton, New Brunswick

N. Kruper
Environmental Protection Service
Edmonton, Alberta

M. Linssen
Pacific Environmental Science Centre
North Vancouver, British Columbia

L. Porebski
Marine Environment Branch
Gatineau, Quebec

J. Princz
Environmental Technology Centre
Ottawa, Ontario

G. Schroeder
Pacific Environmental Science Centre
North Vancouver, British Columbia

R. Scroggins
Environmental Technology Centre
Ottawa, Ontario

T. Steeves
Atlantic Environmental Science Centre
Moncton, New Brunswick

D. Taillefer
Marine Environment Branch
Gatineau, Quebec

L. Taylor
Environmental Technology Centre
Ottawa, Ontario

S. Trottier
Centre St. Laurent
Montreal, Quebec

G. van Aggelen
Pacific Environmental Science Centre
North Vancouver, British Columbia

L. Van der Vliet
Environmental Technology Centre
Ottawa, Ontario

B. Walker
Centre St. Laurent
Montreal, Quebec

P. Wells
Environmental Conservation Service
Dartmouth, Nova Scotia

Federal, Fisheries & Oceans Canada

R. Roy
Institut Maurice Lamontagne
Mont-Joli, Quebec

Federal, Natural Resources Canada

M. Schwartz
Mineral Sciences Laboratory, CANMET
Ottawa, Ontario

B. Vigneault
Mineral Sciences Laboratory, CANMET
Ottawa, Ontario

Provincial

C. Bastien
Ministère de l’Environnement du Quebec
Ste. Foy, Quebec

B. Bayer
Manitoba Environment
Winnipeg, Manitoba

K. Hunter
Ontario Ministry of Environment
Rexdale, Ontario

D. Poirier
Ontario Ministry of Environment
Rexdale, Ontario

J. Schroeder (Chairperson)
Ontario Ministry of Environment
Toronto, Ontario

T. Watson-Leung
Ontario Ministry of Environment
Rexdale, Ontario

Appendix B: Environment Canada, Environmental Protection Service, Regional and Headquarters Offices

Headquarters
351 St. Joseph Boulevard
Place Vincent Massey
Gatineau, Quebec
K1A 0H3

Atlantic Region
15th Floor, Queen Square
45 Alderney Drive
Dartmouth, Nova Scotia
B2Y 2N6

Quebec Region
8^th Floor
105 McGill Street
Montreal, Quebec
H2Y 2E7

Ontario Region
4905 Dufferin St., 2^nd Floor
Downsview, Ontario
M3H 5T4

Western and Northern Region
Room 210, Twin Atria No. 2
4999 - 98th Avenue
Edmonton, Alberta
T6B 2X3

Pacific and Yukon Region
401 Burrard Street
Vancouver, British Columbia
V6C 3S5

Appendix C: Procedural Variations for Culturing Lemna spp. and for Undertaking Growth Inhibition Tests Using Lemna spp., as Described in Canadian, American, and European Methodology Documents

Source documents are listed chronologically by originating agency in the following order: (1) major committees and government agencies, and (2) major authors.

ITM, 1990 represents the Institutet för tillämpad miljöforskning (ITM). This publication gives culturing and toxicity test procedures for Lemna minor compiled and used by the Swedish National Environmental Protection Board in collaboration with the National Chemicals Inspectorate (Institutet för tillämpad miljöforskning), Solna, Sweden.

ASTM, 1991 is the standard guide published by the American Society for Testing and Materials (ASTM) for conducting static toxicity tests with Lemna gibba G3.

APHA, 1992 represents the American Public Health Association (APHA), the American Water Works Association, and the Water Environment Federation, 1992. The publication (in Standard Methods for the Examination of Water and Wastewater - 18th ed.) gives culturing and testing procedures for L. minor which was included as a monitoring tool under the Environmental Effects Monitoring component of the Canadian Federal Pulp and Paper Effluent Regulations. This guideline document was revised in 1996.

USEPA, 1992 is the standard guide published by the Office of Pollution Prevention and Toxics (OPPT), United States Environmental Protection Agency (USEPA), for conducting toxicity tests using L. gibba G3 to develop data on the phytotoxicity of chemicals [under the Toxic Substances Control Act (TSCA)]. It appeared in Title 40, Chapter I, Subchapter R of the Code of Federal Regulations. This guideline document was revised, harmonized with other publications, and re- published (draft) in 1996 (see following citation).

USEPA, 1996 is the draft (April, 1996) standard guideline (OPPTS 850.4400) developed by the Office of Pollution Prevention and Toxics (OPPT), United States Environmental Protection Agency, for conducting toxicity tests using L. gibba G3 and L. minor to develop data on the phytotoxicity of chemicals [under the Toxic Substances Control Act (TSCA), and Federal Insecticide, Fungicide and Rodenticide Act (FIFRA)]. This guideline blends testing guidance and requirements that existed in OPPT and appeared in Title 40, Chapter I, Subchapter R of the Code of Federal Regulations (CFR); the Office of Pesticide Programs (OPP) which appeared in the publications of the National Technical Information Service (NTIS) and the guidelines published by the Organization for Economic Cooperation and Development (OECD). It represents the harmonization of two documents: 40 CFR 797.1160 Lemna Acute Toxicity Test, and OPP 122-2 Growth and Reproduction of Aquatic Plants (Tier I) and 123-2 Growth and Reproduction of Aquatic Plants (Tier 2) (Pesticide Assessment Guidelines, Subdivision J-- Hazard Evaluation; Nontarget Plants) EPA report 540/09-82-020, 1982.

AFNOR, 1996 is the standard guide published by the Association française de normalisation (AFNOR) (test method XP T 90-337,1996). This document gives culturing and toxicity test procedures using L. minor.

OECD, 1998 is the draft (June, 1998) standard procedure published by the Organization for Economic Cooperation and Development (OECD). The guideline is designed to assess the toxicity of substances to L. gibba and L. minor and is based on existing guidelines and standards published by ASTM (1991), USEPA (1996), AFNOR (1996), and the Swedish Standards Institute (SIS) (1995).

SRC, 1997 is the (unpublished) standard operating procedures developed in 1997 by H. Peterson and M. Moody of the Saskatchewan Research Council, Water Quality Section Laboratory, for culturing and testing L. minor. It is based on research conducted by Peterson and Moody (1994-1997) and is a modification of the APHA, 1995-8211 Duckweed (proposed) toxicity test procedure.

DFO, 1979 represents Lockhart and Blouw, 1979. This method, published in a document entitled Toxicity Tests for Freshwater Organisms, E. Scherer (ed.), describes procedures for testing herbicides and sediments with L. minor.

B & P, 1981 represents Bishop and Perry, 1981. This publication describes a standard flow-through growth inhibition test for L. minor. It also compares the relative sensitivity of duckweeds with that of fish and invertebrate species for various test materials.

C & M, 1989 represents Cowgill and Milazzo, 1989. This publication develops rearing conditions and a successful long-term culture medium for maintaining L. gibba G3 and several clones of L. minor. A number of endpoints are examined and compared, and the relative sensitivity of the two duckweed species and various clones to various test materials is investigated.

T & N-K, 1990 represents Taraldsen and Norberg-King, 1990. This publication describes a method for culturing and testing L. minor, primarily for testing effluents. The relative sensitivity of duckweed, Ceriodaphnia dubia, and fathead minnows (Pimephales promelas) to various chemicals and effluents is also discussed.

Appendix D: Review of Culture and Test Media Used in Lemna spp. Growth Inhibition Tests, as Described in Canadian, American, and European Methodology Documents

Source documents are listed chronologically by originating agency.

ITM, 1990 represents the Institutet för tillämpad miljöforskning. This publication gives culturing and toxicity test procedures for Lemna minor compiled and used by the Swedish National Protection Environmental Board in collaboration with the National Chemicals Inspectorate (Institutet för tillämpad miljöforskning), Solna, Sweden.

ASTM, 1991 is the standard guide published by the American Society for Testing and Materials for conducting static toxicity tests with Lemna gibba G3.

APHA, 1992 represents the American Public Health Association, the American Water Works Association, and the Water Environment Federation, 1992. The publication (in Standard Methods for the Examination of Water and Wastewater - 18th ed.) gives culturing and testing procedures for L. minor which was included as a monitoring tool under the Environmental Effects Monitoring component of the Canadian Federal Pulp and Paper Effluent Regulations. This guideline document was revised in 1996.

USEPA, 1992 is the standard guide published by the Office of Pollution Prevention and Toxics (OPPT), United States Environmental Protection Agency, for conducting toxicity tests using L. gibba G3 to develop data on the phytotoxicity of chemicals [under the Toxic Substances Control Act (TSCA)]. It appeared in Title 40, Chapter I, Subchapter R of the Code of Federal Regulations. This guideline document was revised, harmonized with other publications, and re-published (draft) in 1996 (see following citation).

USEPA, 1996 is the draft (April, 1996) standard guideline (OPPTS 850.4400) developed by the Office of Pollution Prevention and Toxics (OPPT), United States Environmental Protection Agency, for conducting toxicity tests using L. gibba G3 and L. minor to develop data on the phytotoxicity of chemicals [under the Toxic Substances Control Act (TSCA), and Federal Insecticide, Fungicide and Rodenticide Act (FIFRA)]. This guideline blends testing guidance and requirements that existed in OPPT and appeared in Title 40, Chapter I, Subchapter R of the Code of Federal Regulations (CFR); the Office of Pesticide Programs (OPP) that appeared in the publications of the National Technical Information Service (NTIS) and the guidelines published by the Organization for Economic Cooperation and Development (OECD). It represents the harmonization of two documents: 40 CFR 797.1160 Lemna Acute Toxicity Test, and OPP 122-2 Growth and Reproduction of Aquatic Plants (Tier I), and 123-2 Growth and Reproduction of Aquatic Plants (Tier 2) (Pesticide Assessment Guidelines, Subdivision J--Hazard Evaluation; Nontarget Plants) EPA report 540/09-82-020, 1982.

AFNOR, 1996 is the standard guide published by the Association française de normalisation (test method XP T 90-337,1996). This document gives culturing and toxicity test procedures using L. minor.

OECD, 1998 is the draft (June, 1998) standard procedure published by the Organization for Economic Cooperation and Development. The guideline is designed to assess the toxicity of substances to L. gibba and L. minor and is based on existing guidelines and standards published by ASTM (1991), USEPA (1996), AFNOR (1996), and the Swedish Standards Institute (SIS) (1995).

SRC, 1997 is the (unpublished) standard operating procedures developed in 1997 by H. Peterson and M. Moody of the Saskatchewan Research Council, Water Quality Section Laboratory, for culturing and testing L. minor. It is based on research conducted by Peterson and Moody (1994-1997) and is a modification of the APHA, 1995-8211 Duckweed (proposed) toxicity test procedure.

SRC, 2003 is the (unpublished) report prepared by M. Moody of the Saskatchewan Research Council, Water Quality Section Laboratory, describing the development of a modified Hoagland’s E+ medium for culturing L. minor. The modified Hoagland’s E+ medium is based on research conducted by Moody and is a modification of the Hoagland’s E+ medium described in the first edition of Environment Canada’s Lemna minor test method document.

ISO, 2005 is the draft international standard test method for testing the effects of water constituents and wastewater on the growth of L. minor, published by the International Organization for Standardization in Geneva, Switzerland.

DFO, 1979 represents Lockhart and Blouw, 1979. This method, published in a document entitled Toxicity Tests for Freshwater Organisms, E. Scherer (ed.), describes procedures for testing herbicides and sediments with L. minor.

pH Adjustment: pH adjust to 6.5 by addition of NaOH or HCl

Sterilization: Stock solutions are sterilized by use of sterilizing filters (pore diameter 0.2µm) or by autoclaving

pH Adjustment: Adjust the pH to 4.60 with KOH or HCl

Sterilization: Autoclave 20 min at 121°C and 1.1 kg/cm²

pH Adjustment: Adjust the pH to 5.0 ± 0.1 with 0.1N KOH or HCl, after autoclaving

Sterilization: Autoclave 20 min at 121°C and 1.1 kg/cm²

pH Adjustment: Adjust to pH 7.5 ± 0.1 with 0.1N NaOH or HCl

Sterilization: Filter medium through a 0.22µm pore size membrane filter into a sterile container

pH Adjustment: Adjust to pH 7.5-8.0

Sterilization: None

pH Adjustment: Adjust the pH to 5.0 ± 0.2 with 0.1N NaOH^{Table note g.6}

Sterilization: Autoclave

pH Adjustment: Adjust the pH of the culture and test media to 5.5 ± 0.5 with NaOH or HCl^{Table note e.13}

Sterilization: Filtration through 0.22 µm filter

pH Adjustment: Adjust the pH to 6.5 ± 0.2 by addition of NaOH or HCl.

Sterilization: Stock solutions I to V are sterilized by autoclaving (120°C, 15 min.) or by membrane filtration (pore diameter 0.2µm); stock solutions VI (and optional VII) are sterilized by membrane filtration only (i.e., these should not be autoclaved).

pH Adjustment: Adjust to pH 8.30 ± 0.05 immediately before testing

Sterilization: None

pH Adjustment: Adjust the pH to 4.6 ± 0.2 with NaOH or HCl

Sterilization: Autoclave 20 min at 121°C and 124.2 kPa (1.1 kg/cm²)

pH Adjustment: Adjust the pH to 5.5 ± 0.2 with NaOH or HCl, if necessary

Sterilization: Autoclave 20 min at 121°C or filter (0.2µm) for longer shelf life

pH Adjustment: NI

Sterilization: NI

Appendix E: General Description of Lemna minor

Taxonomy and Phyletic Relationships

Lemna minor Linnaeus (Arales:Lemnaceae) is a small, vascular, aquatic macrophyte belonging to the family Lemnaceae. Members of the family Lemnaceae are structurally the simplest and the smallest, flowering plants in the world, likely by reduction from more complex ancestors (Godfrey and Wooten, 1979). Most investigators place Lemnaceae in the order Spathiflorae (Arales), relating them to the Araceae through the water-lettuce Pistia (Hillman, 1961).

Four genera are usually recognized: Spirodela, Lemna, Wolffiella, and Wolffia (Hillman, 1961). The fronds (or thalli) of Spirodela and Lemna are flat, more or less oval, in outline and leaf-like. Spirodela bears two or more thread-like roots on each frond, whereas Lemna has only one. The two genera have been grouped in a tribe (Lemneae) (Hegelmaier - 1895) or subfamily (Lemnoideae) (Lawalrée - 1945) (Hillman, 1961). Spirodela has also been considered a subgenus of Lemna (Hutchison, 1934, in Hillman, 1961). Wolffiella and Wolffia have no roots and have been grouped in a tribe (Wolffieae, Hegelmaier) or subfamily (Wolffioideae, Lawalrée) (Hillman, 1961). Wolffia consists of almost microscopic meal-like bodies, whereas Wolffiella is made up of strap-shaped bodies, occurring singly or radiating from a point (Fassett, 1957).

The taxonomy of Lemna spp. (also known as duckweeds) is difficult, being complicated by the existence of a wide range of phenotypes (OECD, 1998). In 1957, Landolt reported the existence of at least two distinct strains of L. minor in the United States that differed in size and in ability to flower in culture (Hillman, 1961). L. perpusilla and non-gibbous forms of L. gibba might easily be mistaken for L. minor (cf. Mason, 1957 in Hillman, 1961). L. gibba differs from L. minor in that the fronds of L. gibba are broadly elliptic to round, its upper surface often has red blotches, and its lower surface is generally swollen (gibbous). L. perpusilla can be distinguished from L. minor by its wing-like appendages at the base of the root sheath and sometimes by its prominent apical and central papillae which are lacking in L. minor (Hillman, 1961; Godfrey and Wooten, 1979). The lack of overwintering turions (dark green or brownish daughter plants), lack of prominent dorsal papules, and of reddish anthocyanin blotches on the ventral side separate L. minor from another closely related species Lemna turionifera Landolt. Taxonomic descriptions and photographs of many Lemnaceae species can be found on the Internet at Wayne P. Armstrong’s Key to the Lemnaceae of western North America (Palomar College/Oregon State University).

Species Description

L. minor is a small, colonial plant with a single, flat, sub-orbicular to elliptic-obovate, leaf-like frond (discoid stem). Each plant is 2- to 4-mm long and consists of a solitary or, in the case of a colony, several (3 to 5) fronds (Hillman, 1961; ITM, 1990). The frond (or thallus) is a complex structure representing both leaf and stem (Hillman, 1961) with the distal end of the frond being foliar and the proximal end being axial (Arber, 1963). The frond is composed largely of chlorenchymatous cells, separated by large intercellular spaces, which are filled with air or other gases and provide buoyancy (Hillman, 1961).

L. minor fronds are obscurely 3-veined (or 3-nerved) and have a smooth convex or somewhat flattened dorsal surface. Although not prominent (Hillman, 1961; Britton and Brown, 1970), the dorsal surface has a small central papilla and usually, a median line of smaller papillae extending near the apex (Godfrey and Wooten, 1979). The lower surface of the frond is convex (or rarely concave when growing in insufficient light or nutrients) (Godfrey and Wooten, 1979). They are green to lime green, glossy when fresh (Godfrey and Wooten, 1979).

The plant has a single root or rootlet that emanates from a deep root furrow in the centre of the lower surface of each frond (Hillman, 1961). The root arises at the node just beneath the lower epidermis and is usually <0.5 mm in diameter, devoid of vascular tissue, and provided with an obtuse or sub-truncate rootcap (Hillman, 1961; Britton and Brown, 1970). Since the entire lower surface of Lemna fronds can absorb nutrients from the medium, and plants can grow well under conditions which entirely prevent root elongation, the functional importance of the root is difficult to evaluate (Hillman, 1961). It has been suggested (cf. Arber, 1920; in Hillman, 1966) that they serve chiefly as anchors to keep the fronds right side up, and to form the tangled masses that aid in dispersal and protection from water motion (Hillman, 1961).

Distribution and Ecology

L. minor is a cosmopolitan species whose distribution extends nearly worldwide (Godfrey and Wooten, 1979). It is widely distributed throughout North America, except the extreme north and in the Bahamas and, is also found in Europe, Asia, Africa, and Australia (Britton and Brown, 1970). In North America, it is found from Newfoundland to Alaska and south to California, Texas, and Florida (Newmaster et al., 1997). In Canada, its distribution extends as far north as Great Slave Lake in the Northwest Territories; Lake Athabasca in Alberta and Saskatchewan; Churchill, Manitoba; James Bay, Ontario; Côte-Nord and Anticosti Island in Québec; and Newfoundland, New Brunswick, Prince Edward Island, and Nova Scotia (Scoggan, 1978).

Duckweeds inhabit lentic environments from tropical to temperate zones, from fresh water to brackish estuaries, and throughout a wide range of trophic conditions (Hillman and Culley, 1978). They can be found in still or slightly moving water of freshwater ponds, marshes, lakes, and quiet streams. Flourishing growth can be found in nutrient-rich, stagnant marshes, bogs, small ponds, or ditches rich in organic matter. Duckweeds are also found commonly near sewer outlets (ITM, 1990).

Duckweeds form an essential component of the ecosystem in shallow, stagnant waters. They are an integral portion of the food chain, providing food for waterfowl and marsh birds such as coots, black ducks, mallards, teals, wood ducks, buffleheads, and rails, and are occasionally eaten by small mammals such as muskrats and beavers. They also provide food, shelter, shade, and physical support for fish and aquatic invertebrates (Jenner and Janssen-Mommen, 1989; Taraldsen and Norberg-King, 1990; APHA et al., 1992; Newmaster et al., 1997). Under conditions favourable for growth, they can multiply quickly and form a dense mat, dominated by a single species (Wang, 1987; ASTM, 1991) made up of mixed genera and species (Riemer, 1993).

Reproductive Biology

Lemna spp. are fast growing, and reproduce rapidly compared with other vascular and flowering plants (Hillman, 1961; APHA et al., 1992). Reproduction of L. minor is usually vegetative (i.e., asexual). New “daughter” fronds are produced from two pockets on each side of the narrower end of an older “mother” frond, very near the point at which the root arises (Hillman, 1961). This end of the frond is usually designated as “basal” or “proximal” since, in an attached daughter frond, it is the portion closest to the mother. The wider end of the frond is denoted as “distal” (Hillman, 1961). Each daughter frond becomes a mother in turn, usually while still attached to its own mother. Groups of attached fronds are called colonies (Hillman, 1961). In Lemna, daughter fronds are produced alternately from each side, developing earlier in one pocket than in the other. Clones of the same species differ as to which pocket produces the first daughter, but this normally remains constant within a clone (Hillman, 1961).

Flowering (i.e., sexual reproduction) in L. minor is rare and occurs only under changing environmental conditions. Photoperiod and high temperatures have been associated with flowering (Landolt, 1957 in Hillman, 1961). Current knowledge indicates that a frond produces only one flower in its lifetime. The flower arises in or near the same meristematic area that produces daughter fronds (Hillman, 1961). Each flower consists of a single flask-shaped pistil (which matures first) and 1 or 2 stamens (which mature at different rates) (Hillman, 1961; Newmaster et al., 1997). These organs are surrounded during development by a membranous sack-like “spathe” open at the top (Hillman, 1961).

The fruit of L. minor is symmetrical, ovoid or ellipsoid, and wingless, and the seed is deeply and unequally 12- to 15-ribbed, with a prominent protruding hilum (Britton and Brown, 1970; Godfrey and Wooten, 1979).

Appendix F: Axenic Culture Techniques for Lemna (Acreman, 2006)

Various species of Lemna (duckweed), vascular, aquatic macrophytes belonging to the Lemnaceae family, can be grown under axenic conditions in liquid media or on nutrient agar using methods similar to those for plant tissue culture. Axenic cultures are free of any contaminants and are literally "without strangers". Good sterile technique and the proper use of a laminar flow hood are essential for axenic culturing of Lemna. Careful monitoring of the cultures and regular testing for contamination is crucial. A basic rule when working with all axenic cultures is to treat the workspace for manipulation of the cultures as you would a surgical operating area. An axenic culture is valuable and if it becomes contaminated, the contamination is not always easy to eliminate. Always make multiple subcultures of the plants to help ensure that at least one or more of them will remain sterile. Tips provided here should help to reduce the potential for contamination of the cultures.

Maintaining a Clean Laboratory

The culture areas such as benches or shelves on which the sterile cultures are kept should be periodically cleaned with 1% sodium hypochlorite (bleach) solution to keep down the levels of dust mites, bacteria and fungal spores. Vacuum the area before applying the solution to reduce any organic contaminants present as they will reduce the effectiveness of the treatment. The bleach solution should be freshly prepared each time and allowed to remain on the surfaces for at least 20-30 minutes. The shelf life of concentrated bleach solution is about 4-6 months once opened, depending on the exposure to light and high temperature. As an alternate solution, granular calcium hypochlorite may be mixed with water at approximately 10g/L providing 70% available chlorine. The dry powder has the added benefit of extended shelf life; if it is kept dry, cool and in an airtight container, it may be stored up to 10 years with minimal degradation. See Appendix 1 for details of preparation of these solutions.

Laminar Flow Hood: Operation and Maintenance

The use of a laminar flow hood is very important to maintaining axenic cultures and good maintenance procedures are critical to the performance of the hood. Handling axenic cultures without such a hood means risking contamination in the long term. Inexpensive hoods costing in the range of $1000-$3000 are available from Envirco Corporation, 1185 Mt. Aetna Road, Hagerstown, MD 21740, USA, (Tel: 1-800-645-1610).

The most important part of a laminar flow hood is a High Efficiency Particulate Air filter (HEPA). Room air is taken into the unit and passed through a pre-filter to remove gross contaminants (lint, dust etc). The air is then compressed and channeled up behind and through the HEPA filter in a laminar flow fashion. The purified air flows out over the entire work surface in parallel lines at a uniform velocity. The HEPA filter is about 99% efficient in removing bacteria and fungal spores of > 22 microns from the air. HEPA filters should be replaced approximately every 7 years for best performance. Routinely check the filter for cracks or damage by sharp instruments. The flow velocity patterns should also be checked annually by a filter service company professional (e.g. H.E.P.A. Filter Services Inc. Tel: 1(800) 669-0037) for any blocked or damaged areas.

If no testing service is available or your budget cannot accommodate the cost of testing, the hood can also be checked for efficiency by using sterility test agar plates (for description of plate preparation see the section below "Testing Lemna for sterility"). It is good practice to periodically check the hood efficiency using this method in between checks by a filter specialist. Spread the plates across the center of the bench and leave them open for at least 24 hours with the hood running. Note the position of each numbered plate. Close the plates, seal them with a double layer of Parafilm and leave in a warm dark location for at least 5 days to monitor for bacterial or fungal growth. If your test indicates that some areas of the HEPA filter are defective, it is possible to repair the filter by injecting silicone sealant if the damaged areas are small. Large patches will cause some air turbulence in the workspace. Ideally the repairs should be done by a company that specializes in HEPA filtered equipment.

Laminar flow hoods are ideally left on at all times. If this is not possible, an ultra-violet germicidal light should be installed to sterilize all surfaces. The fan blower for the hood should then be turned at least 30 minutes prior to using it, to ensure that all the air in the hood will be sterile.

Ideally, the ultra-violet lamp should be left on when the hood is not in use. If this not practical it should at least be left on overnight, and turned off immediately prior to using the hood. UV light can cause skin and eye burn hazards if used improperly. For safe and reliable use of germicidal lamps follow these recommendations:

• Post warning signs near the lamp.
• Clean the bulb at least every 2 weeks; turn off power and wipe with an alcohol-moistened cloth.
• Factors such as lamp age and poor maintenance can reduce performance. Measure radiation output of the bulb at least twice yearly with a UV meter or replace the bulb when emission declines to 70% of its rated output (after about 1 year of normal use). If no UV meter is available replace the bulb once a year.

The working area of the hood, including the bench top and sides should be cleaned with a surface cleaner such as Bio-Clean, Cidex, Sporocidin (VWR) or Viralex (Canadawide). Ethanol is adequate as a disinfectant to reduce microbes but is not recommended as a sterilizing agent since it is not effective as a fungicide or virucide and will not kill bacterial spores. Alcohol (e.g. ethanol) used in concentrations of less than 90% is more effective because the water added to dilute the alcohol allows better penetration of the bacterial cell walls. Optimal concentration range is between 70% and 80%; contact time should be at least 10 minutes. The cleaning agents are sprayed on the surface and left for the appropriate length of time before being wiped clean with paper towels or lint-free tissues. Clean the working area before and after each use.

Keep the hood free of clutter. A direct, unobstructed path must be maintained between the HEPA filter and the area inside the hood where the culture manipulations are being performed. The air downstream from non-sterile objects (such as solution containers, hands etc.) becomes contaminated from particles blown off these objects. Avoid keeping any large containers in the hood.

Pre-filters should be monitored for dust build-up and washed every 2-3 months, depending on how dusty the work area is. They should be thoroughly dry before re-installation. Some pre-filters are not washable and should be discarded when dusty.

Sterilization of Loops and Other Instruments

Bunsen burners and other continuous flame gas burners are effective but can produce turbulence, which disturbs the protective airflow patterns of the laminar flow cabinet, and additionally, the heat produced by the continuous flame may damage the HEPA filter. If a gas burner must be used, one with a pilot light should be selected and the burner should not be closer than 20 cm from the HEPA filter. Electric sterilizers may also be considered. Alternatively, disposable plastic loops and needles may be used for culture work where electric incinerators or gas flames are not available.

Hand Cleaning

Before performing any manipulations or subculturing, remove any rings and wash hands thoroughly with an antibacterial soap followed by a cleanser e.g. One-Step, Endure or 70% ethanol. Pay attention particularly to the areas of your hands that may come in contact with the culture vessels or transfer loops. Examination gloves (e.g. Nitrile) may be used and sprayed with ethanol before handling cultures.

Preparation and Sterilization of Media

Autoclaving is the most widely used technique for sterilizing culture media, and is the ultimate guarantee of sterility (including the destruction of viruses). A commercial autoclave is best, but pressure cookers of various sizes are also suitable. Sterility requires 15 minutes at a pressure of 15 psi and a temperature of 121°C in the entire volume of the liquid (i.e. longer times for larger volumes of liquid; approximately 25 min for 100-200 mL, 30 min for > 200-1000 mL, 45 min for 1-2 L and 60 min for > 2 L). It is best to autoclave the medium in small batches to minimize the time for effective autoclaving and avoid chemical changes in the medium due to long exposure to high temperatures. Large loads in the autoclave should be avoided, as they will require more time to reach the sterilization temperature and there is the risk that the media may not be properly sterilized.

Heat sensitive indicator tape that changes colour should be used on the outside of media vessels and packages of material for sterilization to indicate that the appropriate temperature has been reached. They are NOT a guarantee of sterility and only indicate that the material has been through the sterilization process. It is important to ensure that large volumes of media or large loads in the autoclave have reached the appropriate temperature for sterilization. Commercially available biological indicators in sealed ampoules (e.g. Raven Biological Laboratories) or chemical integrator strips (e.g. STEAMPlus Steam Sterilization Integrator strips from SPS Medical) may be used. A simple, alternate method is to put a small piece of autoclave tape into a Pasteur pipette, heat-seal the tip and cotton-plug the other end. Attach string to the pipette and lower it into the medium, keeping the plugged end about 10-15 cm above the liquid surface. Tape the other end of the string to the outside of the flask so that you can easily pull the indicator out. Recover the indicator after the run and confirm that it too has changed colour. The latter method is not as reliable as the biological or chemical integrator strips.

Autoclave efficiency should also be regularly checked with biological indicator tests containing bacterial spores. There are commercially available test indicator kits (e.g. VWR Cat #55710-014) that use spores of Bacillus stearothermophilus that are rendered unviable at 250 °F or 121 °C. For the test, spore strips or ampoules of B. stearothermophilus are autoclaved, incubated for 48 hours in Tryptic Soy broth, then observed for any sign of growth, which would indicate that the autoclave is not sterilizing properly.

Bottles and tubes containing media should be no more than 2/3 full to prevent boiling over. If using screw capped media bottles leave the caps slightly unscrewed. Flasks can be loosely plugged with a bung made of non-absorbent cotton wool covered with cheesecloth and with a square “skirt” of either Bio- Shield Wrap (VWR 59100-234) or aluminum foil over the top. After autoclaving, the pressure release valve on the autoclave should not be opened until the temperature has cooled to below 80°C. As the pH of media rises during autoclaving, allow at least one day before using the media in order for the pH to readjust to the level set prior to sterilization.

Autoclaving is a process that may have negative effects on media as components may be broken down on prolonged exposure to heat. Precipitates of phosphate (white) or iron (yellow) may occur at times. To avoid this problem the iron and phosphate solutions can be sterilized separately and added aseptically after autoclaving. Precipitates in media may also be avoided by filter-sterilizing using filters of pore size 0.22 microns or smaller.

Agar plates are convenient for long-term maintenance of Lemna. They are usually prepared at least 2 days before use and allowed to dry in the laminar flow hood before double sealing with Parafilm (VWR) or Duraseal (VWR or Sigma). If plates are not to be used in a week or so after preparation they should be wrapped in plastic film, inverted and stored at room temperature for a few days to monitor for contamination before storing in the refrigerator. For slants place the filled tubes on a 45^o angle and allow agar to gel with the caps slightly unscrewed to prevent excessive condensation build-up. After they are dry, tighten the caps securely and refrigerate after monitoring for contamination at room temperature. Slants and agar plates may be stored for several months at 4^oC.

Transfer Techniques

The following procedures should always be used when transferring cultures:

• All culture vessels, transfer tools, cotton-plugged pipettes and media must be sterilized and ready to use. Media should be at room temperature.
• Loops should be first dipped in 95% Ethanol and then sterilized in a flame or electric sterilizer for 15 seconds until they are red-hot before use. Cool the loop by touching it to sterile agar or liquid before using it to pick up the plants. The flame from a gas burner effectively sterilizes small glass or metal objects, such as inoculating loops, but one must avoid “frying” the plants by contact with objects heated in a flame.
• Clear the laminar flow hood so that nothing is between the path of the airflow coming from the HEPA filter and the area where the subculture is being done. Do not allow anything to come in contact with the HEPA filter.
• Clean the bench of the laminar flow hood thoroughly just before use but avoid spraying any solutions on the HEPA filter.
• Wash hands thoroughly or put on gloves (see above) immediately prior to subculturing.
• Flame all openings of glass culture vessels for 15 seconds before and after transferring the new culture material to them.
• To minimize contamination, always carry out the transfers at least 6 inches (15 cm) from the front of the hood to ensure that the area is not contaminated by room air. Where possible, perform the operation at eye level.
• Don’t touch anything that will come in contact with the culture and if you do touch it, sterilize it again before using it.
• When subculturing to screw-capped tubes, loosen the caps slightly before picking up the plants to be transferred to prevent the plants from falling from the loop while opening tightly sealed tubes.
• Avoid talking, singing, whistling, coughing or sneezing in the direction of things that should be sterile. Long hair, if not tied back, may be a source of contamination.
• Work quickly to minimize the time that the culture vessels are open.
• Try not to touch the edges of the Petri plate covers. Hold the cover by the top.
• Seal all Petri plates with a double layer of Parafilm or Duraseal. Monitor carefully for cracks. (Dust mites are attracted to the smell of the media and may crawl into the sterile plates.)
• Monitor plates every 2-3 days for presence of contaminants.
• Transfer the cultures every 2-3 weeks for best results.

Testing Lemna for Sterility

Contaminants such as bacteria and fungi are readily apparent when Lemna is cultured in a medium enriched with organic components e.g. Hoagland's E+. If the plants are not cultured routinely in such medium they should be periodically tested for sterility by removing a few plants and placing them in Hoagland’s E+, which contains 1% sucrose, 0.6% Bacto-tryptone (or peptone) and 0.1% yeast extract. This can be done in liquid culture or on agar plates. Contamination by fungi and bacteria will usually show up in solutions or on agar plates within several days. If the solution becomes cloudy or colonies of bacteria or fungi grow on the plates you can try the cleaning technique described below or obtain a new axenic culture from an outside source.

Cleaning Lemna Plants

If Lemna plants become contaminated they can be made sterile again but the techniques require time and patience. In order to do this, plants connected in clonal clusters should be separated from each other. Individual plants should be dipped in a 0.5% solution of sodium hypochlorite (10% Clorox® or Purex® bleach solution) for at least one minute. Treat plants with bleach for varying amounts of time to ensure that you have at least one living culture that is sterile. Be sure to rinse the plants in several changes of sterile medium or sterile water before transferring to dilute growth medium (e.g. modified Hoagland’s medium containing 1% sucrose). Examine your plants after rinsing them in fresh medium. Properly sterilized plants will have a small green area in the bud zone along the center of the frond. If there is no green bud remaining, the plant was treated too long and is dead. Since only a small bud is left to re- grow after surface sterilization, it may take some time before sufficient plant material is available to do experiments.

According to Landolt (1987), about 1-10% of the plants normally survives this treatment and becomes axenic. Plants that do survive this sterilization technique (and are not contaminated or infected by fungal molds or bacteria) can be transferred to an enriched medium such as Hoagland’s E+ in liquid form or solidified with 1.25 % Difco-Bacto agar in Petri plates or tubes.

Long Term Preservation of Lemna by Cryopreservation

Cryopreservation is a technology to store living cells at ultra-low temperatures indefinitely. Valuable strains of Lemna can be maintained at ultra-low temperatures in the liquid or vapour phase of liquid nitrogen to preserve their genetics and to maintain the cultures over long terms without maintenance through subculturing (Day 1995; Kartha 1985).

The techniques used must minimize the formation of destructive intracellular ice crystals which damage cell membranes and walls. The basic procedures of cryopreservation involve removal of the free water by osmotic agents followed by addition of cryoprotectants such as sucrose and glycerol. Cultures are stored in cryovials and then may be slowly cooled in a -80^oC freezer to minimize ice crystal formation, followed by immersion directly into liquid nitrogen at -196^o C. Cultures are regenerated by rapid thawing in a water bath at 45^oC and subcultured to fresh medium.

For further information, please refer to the following websites:

Armstrong, Wayne. Treatment of the Lemnaceae. Palomar University

Cross, John. The Charms of Duckweed.

McCauley, D. Aseptic technique. GlobalRPh Inc.

Appendix 1: Solutions for Disinfecting Surfaces

1% sodium hypochlorite solution (0.5 L)

1. Commercially prepared bleach is normally a 5% sodium hypochlorite solution. Prepare the dilution just before use.
2. Use a 500 mL graduated cylinder to measure 100 mL of commercial bleach. Add 400 mL of distilled or deionized water to dilute the bleach in the graduated cylinder to a volume of 500 mL.

Chlorinated solution from powder

1. Add 10 g of granular calcium hypochlorite to 1 liter of distilled water.
2. Stir vigorously and allow the mixture stand for 6 hours or overnight. Wear gloves and mask as chlorine gas is corrosive. If possible, make the solution in a fume hood.
3. Filter the supernatant into a clean plastic jug and stopper tightly. If storing in glass the solution should be kept in the dark.

70% ethanol (used to wipe down laminar flow hood surfaces and to spray gloves)

1. Use a 500 mL graduated cylinder to measure 370 mL of 95% ethanol.
2. Add distilled water to bring the volume of liquid in the cylinder to 500 mL.
3. Keep in a tightly capped container.

Appendix G: Logarithmic Series of Concentrations Suitable for Toxicity Tests^Footnote 77

Appendix H: Biological Test Methods and Supporting Guidance Documents Published by Environment Canada’s Method Development & Applications Section^Footnote 79