This project has received funding from the European Union’s Horizon 2020 research

and innovation programme under grant agreement No 731065

Project Title: AQUACOSM: Network of Leading European AQUAtic MesoCOSM Facilities Connecting Mountains to Oceans from the Arctic to the Mediterranean

Project number: 731065

Project Acronym: AQUACOSM

Proposal full title: Network of Leading European AQUAtic MesoCOSM Facilities

Connecting Mountains to Oceans from the Arctic to the Mediterranean

Type: Research and innovation actions

Work program topics addressed:

H2020-INFRAIA-2016-2017: Integrating and opening research infrastructures of European interest

Deliverable No 4.1.2: Standard Operating Protocol (SOP) on Sampling and Analysis of Zooplankton

Due date of deliverable:

Actual submission date:

Version: V1.0

Main Authors: Deniz Başoğlu, Meryem Beklioğlu, Robert Ptacnik, Lisette de Senerpont Domis, Marko Reinikainen, Jens Nejstgaard, Gérard Lacroix

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Project ref. number 731065

Project title

AQUACOSM: NETWORK OF LEADING EUROPEAN AQUATIC MESOCOSM FACILITIES CONNECTING MOUNTAINS TO OCEANS FROM THE ARCTIC TO THE MEDITERRANEAN

Deliverable title Standard Operating Protocol (SOP) on Sampling and Analysis of Zooplankton

Deliverable number D4.1.2

Deliverable version V1.0

Contractual date of delivery

Actual date of delivery

Document status

Document version V1.0

Online access Yes

Diffusion Public

Nature of deliverable Report

Work package WP4.1

Partner responsible METU, WCL, …

Author(s) Deniz Başoğlu, Meryem Beklioğlu, Robert Ptacnik, Lisette de Senerpont Domis, Marko Reinikainen, Jens Nejstgaard, Gérard Lacroix

Editor Deniz Başoğlu, Meryem Beklioğlu, Robert Ptacnik, Lisette de Senerpont Domis.

Approved by Jens Nejstgaard (FvB-IGB)

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EC Project Officer Agnès Robin

Abstract This deliverable is a Standard Operating Protocol (SOP) that describes the methods for sampling and analysis of mesozooplankton from mesocosm experiments carried out in all aquatic environments (fresh and marine waters). It gathers best practice advice with a focus on sampling, counting and other analyses of mesozooplankton as well as Quality Assurance/Quality Control (QA/QC) practices.

Use of this SOP will ensure consistency and compliance in collecting and processing mesozooplankton data from mesocosm experiments across the AQUACOSM community, in Europe and beyond.

Keywords Zooplankton Analysis, Mesozooplankton, Sampling, Enumeration, Standard Operating Protocol, Freshwater, Brackish, Marine, Mesocosm

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Table of Contents

1. Executive summary 6

2. Definitions and Terms 7

3. Cross References 8

4. Materials and Reagents 8

5. Health and Safety Indications 9

5.1 General Information 9

5.2 Safety Instructions for Sampling 10

5.3 Working and Personal Protection 10

5.4 Use, storage and disposal of reagents and chemicals 10

5.5 Use, storage and disposal of equipment 11

6. Environmental Indications 11

7. Sampling Zooplankton 12

7.1 Prior to sample collection 12

7.2 Required Strategies 12

7.3 Preparation and calibration of sampling equipment 12

7.3.1 Overview of equipment and instruments 12

7.3.2 Sample bottle preparation and preservation 12

7.4 Sampling Equipment 13

7.4.1 Plankton Nets 13

7.4.2 Sampling with plankton nets: 15

7.4.3 Using water samplers to collect zooplankton in mesocosms 16

7.4.4 Additional materials generally needed for zooplankton sampling 18

7.4.5 Sampling Containers and Sample Bottles 18

7.5 Sampling Design for Shallow and Deep Mesocosms 19

7.5.1 Sampling time and frequency 19

7.5.2 Shallow mesocosms 19

7.5.3 Deep mesocosms 20

7.5.4 Best Practice Advices on selecting the right sampling instrument 21

7.6 Best practice advice on preservation and storage 22

8. Quantitative Analysis of Zooplankton 23

8.1 Sample Preparation 23

8.1.1 Subsampling 23

8.2 Counting Procedure (Enumeration) 25

8.3 Best Practice Advice on subsampling and counting 26

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8.4 Taxonomy and nomenclature 27

9. Estimating the biomass 28

9.1 Crustacean Zooplankton 28

9.1.1 Measuring the length or dimensions of an organism 28

9.1.2 Length-weight relationships (Adopted from Bottrell et al. (1976) [48]) 28

9.1.3 Estimating the dry weight (Adopted from [49]) 29

9.2 Rotifers 29

9.2.1 Predicting dry weight using geometrical formulae (Adopted from [49]) 29

9.3 Estimating Mass as Carbon Content (Adopted from [49]) 30

9.4 Estimating biomass: other methods in literature 30

10. Size Distribution 31

10.1 Size Diversity 31

10.2 Normalised biomass-size spectrum (NSS) 32

11. Quality assurance and quality control 33

12. Appendix 34

Appendix A 34

Appendix B 36

Appendix C 36

Appendix D 38

Appendix E 40

13. References 41

14. Checklist for the next version 48

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1. Executive summary

This Standard Operating Procedure (SOP) describes how to sample, identify and determine composition, abundance and biovolume of meso- and macrozooplankton from mesocosm experiments carried out in all aquatic environments (fresh, brackish and marine waters). It gathers best practice advice with a focus on sampling, counting and other analyses of mesozooplankton as well as Quality Assurance/ Quality Control (QA/QC) practices. It covers guidance on health, safety and environmental information, best practice advice on materials and methodology and QA/QC procedures to be followed during the sampling, analysis and counting of meso/macrozooplankton samples. It is designed to be compliant with the European Standard EN 15110:2006 [1]. Use of this SOP will ensure consistency and compliance in collecting and processing mesozooplankton data from mesocosm experiments across the AQUACOSM community, in Europe and beyond. Zooplankton are heterotrophic organisms living in open water whose distribution is primarily determined by water currents and mixing. The size of zooplankton ranges from a few microns to 20 µm (nanozooplankton mainly protozoans), 20-200 µm (microzooplankton, large protozoans and small metazoans), 200-2000 µm (mesozooplankton), and >2 mm (macrozooplankton) whereof those >20 mm are often distinguished as megazooplankton. This SOP focus on zooplankton >200 µm, as they are traditionally sampled with different methods than micro- and nanozooplankton and generally dominate the zooplankton biomass in mesocosm experiments. The dominant groups in mesocosm experiments in most water types are meso- and macrozooplankton such as crustaceas (dominated by copepods and cladocerans) often followed by rotifers [2]. In marine systems notable metazoan zooplankton normally also include Larvaceans and Chaetognaths.

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2. Definitions and Terms

Biomass the amount of living matter present in the mesozooplankton sample [3]

Epilimnion or

Upper mixed layer

water above the pycnocline, i.e. thermocline (freshwater and marine systems) or halocline (marine) a in a stratified body of water [1]

Fixation protection from disintegration of the morphological structure of organisms [1]

Halocline vertical zone in the oceanic water column in which salinity changes rapidly with depth [4]

Hypolimnion or deeper layers

water below the pycnocline, i.e. thermocline (limn./mar) or halocline (mar) a in a stratified body of water [1]

Littoral zone shallow marginal zone of a body of water within which light penetrates to the bottom; usually colonised by rooted vegetation [1]

Pelagic zone body of water beyond the littoral zone [1]

Plankton organisms drifting or suspended in water, consisting chiefly of minute plants or animals, but including larger forms having only weak powers of locomotion [1]

Sampling site (Sampling station)

general area within a body of water from which samples are taken [1]

Stratified water freshwater (generally lakes/standing water) or marine waters with a strong density gradient (normally temperature and/or in marine systems salinity) resulting in an upper, normally warmer, mixed/isothermal layer floating on a denser, usually colder and or more saline, also isothermal water

Subsampling collection of a sub-sample that consists of a known fraction of the total sample and that is representative of the quantity and species composition of the latter [1]

Thermocline (metalimnion)

layer in a thermally stratified body of water in which the temperature gradient is at a maximum [1]

Zooplankton animals (heterotrophic organisms) present in plankton [1]

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3. Cross References

The SOPs that will be provided by AQUACOSM will be listed here in the following versions when the different SOPs are completed. The SOPs that will be provided by AQUACOSM will be for:

1. Phytoplankton (Deliverable 4.1.1) 2. Zooplankton (this SOP) 3. Periphyton (Phytobenthos) (Deliverable 4.1.3) 4. Water Chemistry (Physical and Chemical Elements of Water) (Deliverable 4.1.4) 5. High-Frequency Data Collection (Deliverable 4.1.5) 6. QA/QC (Deliverable 4.1.6) 7. Data Management (Deliverable 4.1.7)

A general description for water sampling is covered under the Water Chemistry SOP.

4. Materials and Reagents

Different preserving solutions with different areas of application are available in the literature. Some of the preserving reagents are summarized in Table 4-1.

Table 4-1 The materials and reagents used in analysis of mesozooplankton

Name and concentration Composition Storage

Ethanol (normally 70-99 %) [1] Ethanol, water

Instead of pure ethanol, cheaper methylated spirit can be used. If mesozooplankton is stored for molecular analysis, a solution from pure ethanol should be used (70%).

Solvent cabinet

Formaldehyde (37 % by volume) [1] See Appendix A for detailed information about preparing formaldehyde solution.

Fume Cabinet/Hood

Buffered sucrose formaldehyde (Buffered Formalin) [5]

See Appendix A for detailed information about preparing buffered formalin solution.

Acidified Lugol’s Iodine See Appendix A for detailed information about neutral, acidic and alkaline Lugol’s solution.

Fume Cabinet/Hood

In addition, different narcotizing agents such as Tricaine Methanesulfonate (MS-222), magnesium chloride, carbonated water, chloroform and methyl alcohol are commonly used in marine sampling [6].

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According to [1], preserving solutions should meet the following requirements:

● The most frequently used preservatives in mesozooplankton research used to be Lugol’s Iodine in freshwater and formaldehyde in both freshwater and marine systems, while ethanol is presently increasingly used for fixation, because it presents less health hazards and allows genetic analysis. One should note that formaldehyde can only be used with ‘special permission’ in some laboratories as it is associated with allergies and cancers. For that reason, it should be handled with care and authorized personal should be contacted for permission and guidance before use.

● The effect of the fixative (concentration, method of addition) on the discoloration, deformation and even loss of organisms by chemical shock or otherwise, should be considered before choosing fixative. Alternatively, with increasing use of live analysis of zooplankton – this can be evaluated to avoid fixation altogether. This would also allow other chemical analyses of the same organisms down-stream. This will be further discussed in the next version of this SOP.

● The preservative should effectively prevent the microbial degradation of organic matter for at least the storage period of the samples.

● The preservative should guarantee a good recognition of taxa for at least the storage period of the samples.

● The preserving solutions should be kept in closed bottles, which should be stored in a fluid tight plastic box or container during transportation to and from the sampling (to allow fixation directly after sampling and to prevent spillage of fixative. Use of gloves and pipettes to transfer the solution to the plankton samples are recommended [1]. Most fixatives, such as Lugol’s Iodine and formaline need to be renewed after certain time in storage, typically due to oxidation. Oxidation rate depends on fixative, buffer and storing conditions. Samples should be stored dark and cold (but above freezing) for best preservation.

● The advantages and disadvantages of each of these solutions are detailed in Appendix A. It should be noted that the reagent may affect the size of the organisms (shrinkage). The shrinkage may differ depending on fixative concentration and status of the fixed organisms before fixation. Please see Appendix A for more information.

5. Health and Safety Indications

5.1 General Information

In this section, general guidance on the protection of health and safety while sampling, analysing and counting mesozooplankton from mesocosm experiments will be provided to minimize the risk of health impacts, injuries and maximize safety. The users of this SOP are expected to be familiar with the Good Laboratory Practice (GLP) of World Health Organization (WHO) [7] and Principles on GLP of Organisation for Economic Co-operation and Development (OECD) [8]. Health and Safety Instructions of the mesocosm facility, if there are any, shall be followed properly to protect the people from hazardous substances and the harmful effects of them. According to preventive employment protection measures to avoid accidents and occupational diseases (on-site or in the laboratory), the work should be practiced consistent with national and EU regulations (see the OSH Framework Directive 89/391/EEC, [9]). Other regulations and guidelines can be found in EU – OSHA website (European directives on safety and health at work [10]). All necessary safety and protective measures shall be taken by the users of this SOP and the scientist-in-charge shall ensure that those measures comply with the legal requirements.

The table below summarizes the hazards and risks and the measures for preventing them associated to the laboratory studies carried out on mesozooplankton:

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Table 5-1 Hazards and risks associated with laboratory work

Occupations at risk

Hazards/Risks Preventive Measures

Laboratories o Risk of inhaling chemicals that are used for mesozooplankton sample fixing

o Accidental spills

o Use of fume cabinets/hood o Safe handling and transport of samples o Appropriate personal protection (protective

gloves) and hygiene measures o Storage of samples in glass bottles to

minimize diffusion of the fixative o Safe handling and transport of samples

o Use of closable cooling boxes to transport samples between field and laboratory

o Using of closable plastic boxes to store the sample bottles

5.2 Safety Instructions for Sampling

Please see the Water Chemistry SOP (Deliverable 4.1.4) for safety instructions for sampling.

As a best practice advice, if the samples are collected from a boat, always have a shore-based contact in case of emergencies. Check the weather forecast to ensure safe and effective working conditions. For safety reasons, it is recommended that field work should not be undertaken by unaccompanied persons, but by a minimum of two people [1].

5.3 Working and Personal Protection

Please see the Water Chemistry SOP (Deliverable 4.1.4) working and personal protection equipment suggested for use in water sampling.

5.4 Use, storage and disposal of reagents and chemicals

The user of this SOP needs to visit the Safety Data Sheets (SDSs), which is provided by the manufacturer for any chemicals. The SDSs contain necessary information for protection before using, storing and disposing the reagents as well as other chemicals. Only experienced personnel should be responsible for the use of reagents, preservatives and chemicals in a way compatible with the laboratory rules. Important: Use of formaldehyde as a fixative can result in “the generation of bis(chloromethyl)ether” - that is a potential human carcinogen [11]. In addition, formaldehyde, is an irritant and can enter the body via inhalation and damage tissues seriously. For more information on formaldehyde, see the substance information [12] provided by European Chemicals Agency (ECHA).

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5.5 Use, storage and disposal of equipment

Zooplankton sampling equipment needs to be cleaned in between sampling events [13]. The storage of equipment should be compatible with the operating manual/instructions for use. Cleaning, disinfection and storage of equipment should be taken care under the risk of spreading parasites, diseases, foreign species, and pathogens.

6. Environmental Indications

A plan for the disposal of mesocosm waste needs to be prepared prior to the experiments. The plan must comply with the EU Waste Legislation [14] and The List of Hazardous Wastes [15] provided by the European Commission. The Safety Data Sheets (SDSs) need to be followed for the disposal of reagents and chemicals prior to waste disposal.

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7. Sampling Zooplankton

7.1 Prior to sample collection

The AQUACOSM Standard Operating Procedure (SOP) on sampling, analysis and counting of mesozooplankton needs to be reviewed by the research team prior to initiating the sampling. The sampling method to be employed, sample type and volume, equipment and supplies needed for sample collection must be identified prior to the start of a sampling event based on the objectives and main questions of the study. In addition, the type of mesozooplankton organisms such as larger crustaceans or smaller ones like rotifers, and larval meroplankton may require different methods for sampling, counting and analysis [2].

7.2 Required Strategies

As suggested in [1] and [5], the documentation that should be available before the start of field work is listed below:

● Sampling plan; including the description of objectives and strategy and methods, data QA/QC objectives – (See SOP on QA/QC, Deliverable 4.1.6)

● Applicable safety instructions ● The registration (metadata) forms (provided in Appendix B).

7.3 Preparation and calibration of sampling equipment

The sampling equipment should be checked and prepared in time before the sampling and left in proper order for the next sampling. The required actions to prepare the equipment for mesozooplankton sampling are summarized from [1] as follows:

7.3.1 Overview of equipment and instruments

● Nets, especially finer meshed (< 200 µm) should be washed “in warm freshwater with detergent or in an ultrasonic water-bath”, to minimize the risk of clogging and to ensure optimum filtration capacity [1], especially after waters with high concentrations of large diatoms or other organisms that can stick to the mesh and permanently clog the net if dried.

● Before use, plankton nets and the draining cups should be visually checked for tears or holes. ● The string (rope) should be checked to ensure that it is securely attached to the sampling equipment

(plankton net/volume sampler). It should be visually checked for the depth marking on the line or the winch.

● The closing mechanism of the sampler should be checked to ensure that it is well functioning. ● To prevent cross-contamination of organisms between mesocosms, the sampling equipment should be

cleaned thoroughly between uses in the different waters.

7.3.2 Sample bottle preparation and preservation

In order to ensure minimal change of the sample content due to e.g. predation of smaller organisms in the concentrated sample (codend feeding) it is suggested to fix the samples as soon as possible after sampling. This can either be done by adding fixative to the sample bottles before commencing the fieldwork. If it is not possible to add fixatives to the bottles (e.g. due to restrictions of use of toxic chemicals at the site), an alternative is to add non-toxic narcotizers to the bottle to prevent condend feeding before fixing. It is also suggested to mark the

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bottles before sampling (see Appendix X for details on labelling). To ensure safe marking, it is suggested to add a water-resistant paper with sample identification written with a pencil inside the bottles.

For more information on the effectiveness of the devices for specific type of mesozooplankton, one can visit Welch (1948) [16], Edmondson and Winberg (1971) [17], de Bernardi (1984) [18], DeVries and Stein (1991) [19], Harris et al. (2000) [20], Schwoerbel (2016) [21], Manickam et. al. (2019) [22] and Santhanam et. al. (2019) [23].

7.4 Sampling Equipment

Zooplankton sampling from mesocosms can be performed for qualitative and quantitative analysis. Qualitative mesozooplankton sampling mainly provides information about “the species composition, number of species, size distribution and relative dominance of species and groups of mesozooplankton” [1]. On the other hand, quantitative sampling provides information on the quantity of mesozooplankton per unit volume of water, as well as a basis to calculate the total biomass for the mesozooplankton and individual mesozooplankton species [1]. In addition, size diversity can be estimated based on quantitative analysis carried out. Whenever there are no significant difference in effort of sampling, it is of the latter reasons recommended to conduct fully quantitative sampling. There are many different devices to sample mesozooplankton; including bottles, traps, tubes, pumps and nets. The effectiveness of these different techniques varies according to the type of mesozooplankton [2], type of the mesocosm (depth of strata), type of the samples (point or integrated), volume of water to be sampled and the studied problem [18].

7.4.1 Plankton Nets

Plankton nets, have been the most common tool to sample zooplankton for soon 200 years. They are available in various dimensions and mesh sizes that vary according to the study objectives (e.g. Figure 7-1). However, drawbacks with plankton nets is that they suffer from variable rate of under-sampling due to the resistance for water to pass through the meshes, and thus “loss of organisms through the meshes, net avoidance, and variation in filtration efficiency” [6]. The degree of under-sampling generally increases with decreasing mesh size and increasing amounts of objects caught by the net. “Net efficiency is affected by a series of factors including characteristics of the fabric used to construct the net (i.e. the gauze), mesh sizes, porosity, speed of sampling, avoidance by target organisms, escape of sampled organisms, and clogging” [18]. Moreover, net sampling only measure the average density of organisms integrated over a certain volume but cannot measure the density or distribution of organisms on small spatial scales [6].

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In order to optimize the efficiency of water filtering, plankton nets should have a large filtering surface relative to their opening. This can be achieved by a long net compared to the opening (i.e., opening diameter of 30 cm and length of about one meter), as recommended by [1]. A larger filter surface can also be achieved with a

shorter net by using a cylindrical net section above the conical part in comparison with a conical plankton net with the same opening diameter and length [1] (compare Figure 7-1a-c), and may also be achieved by decreasing the opening compared to the net surface area as with an Apstein-cone (Figure 7-1d)

Figure 7-1. Different types of plankton nets: a) definition of net length and opening diameter and demonstrating how the filtering surface is increased by a cylindrical net section above the conical part, b) conical plankton net, c) cylindrical plankton net (Indian Ocean Standard Net), and d) a net with Apstein cone (Adopted from [1] and [24]).

According to the size classes of the mesozooplankton, appropriate mesh sizes can be selected. The appropriate mesh size for the plankton nets to be used qualitative sampling are summarized in Table 7 – 1 below. For higher sampling efficiency, (i.e. a balance between catching small sized organisms with limited escape ability, and faster swimming larger sized plankton), use of different nets with different mesh sizes and hauling speed, is recommended. NOTE that the size of most mesocosms do not allow substantial sampling of larger meso- and macrozooplankton as their abundances are too low to allow removal in numbers adequate for statistical analyses without exhausting the content of the mesocosms. In such cases it is recommended to sample these by emptying the mesocosms through a net at the end of the experiment.

a b c d

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Table 7 – 1: Appropriate mesh size for qualitative sampling of different sizes of mesozooplankton

Targeted plankton Size (µm) Mesh Sizea Notes on the nets with specified mesh sizes

Microzoo-plankton

20 - 200 Typically 20-90 µm, in Freshwater 90 µm nets with Apstein collars are commonly used for freshwater crustaceans, and 40-50 µm to include Rotifers.

+ Useful in oligotrophic waters

- Are expected to have clogging problems, that can be partially ameliorated with an Apstein-collar. 90 µm is a common compromise between filtering efficiency and small mesh size.

Mesozooplankton 200-2000 200 μm (or 150 - 180 µm) mesh is used to catch all mesozooplankton, including predatory species in freshwater

+ Useful in sampling rotifers and crustaceans larger than 200 µm, but many rotifers and early crustacean development stages are technically microzooplankton (<200 µm) and will be strongly under-sampled.

- Low efficiency in sampling both types 500-2000 300 - 350 µm

Macrozooplankton >2000 1000 µm or larger, with non-filtering codend for delicate macrozoopankton, such as medusae.

Slow swimming macrozoopankton such as medusae can be collected with e.g. 1000 µm mesh nets, often with large non-filtering codends to minimize damage. However, fast swimming “zooplankton” like krill is caught with larger and faster devises like Isaak-Kidd trawls. But such animals are not generally possible to hold in mesocosms.

a Mesh sizes mentioned should be regarded as for guidance. Mesh sizes will vary somewhat from manufacturer to manufacturer.

7.4.2 Sampling with plankton nets:

Nets and the buckets to be used for sampling should be cleaned, checked for holes and tears prior to sampling. Then, the collection bucket is attached to the cod end of the net. The bridled end of the net should be attached to a string (rope) with markings for determined depths, to measure the sampling depth.

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The net should be retrieved by pulling it back to the surface with a steady constant speed, ca 0.5 m/s or slower if clogging is severe for smaller meshed nets. When at the surface, the catch can be rinsed into the codend by dipping the net and rapidly lifting it out of the water (keeping the opening above water at all times).

When depth stratified samples are needed nets with closing mechanisms may be used, such as Apstein closing nets [24], or a Clarke-Bumpus sampler [25] that is a plankton net connected to a flowmeter, allowing measurement of the volume of water filtered by the sampler. For additional information and best practice advices on using Clarke-Bumpus plankton samplers, please refer to [18] and [26].

Then, the contents of the plankton net should be rinsed to the collection bucket by spraying water against the outside of the net. The collection bucket should be removed from the net vertically. The contents in the collection bucket can be concentrated by swirling the bucket.

Risk of contamination through net sampling: Plankton nets represent large surfaces and are therefore prone to cause contamination among mesocosms and samples. Proper rinsing after every single sample is mandatory in order not to contaminate subsequent samples and mesocosms (i.e. animals that are not washed from the net after a net haul may end up in the mesocosm sampled/in the sample form the mesocosm sampled next). Contamination is particularly problematic if mesocosms contain different communities, which is an issue in experiments involving manipulation of connectivity. Note that protists and bacteria will easily be transferred among mesocosms through net sampling. To prevent cross-contamination of organisms between mesocosms, the sampling equipment should be cleaned thoroughly between uses in the different waters. Moreover, in some mesocosms, there are sampling ports to avoid contamination among mesocosms.

Using plankton nets for quantitative sampling, one shall measure and keep record of the quantity of water, that passes through the net. The volume of filtered water should be calculated with the following formulation, as given in [18]:

𝑉 = 𝜋 × 𝑟2 × 𝑑

where V is the volume of water filtered by the plankton net, r is the radius of the mouth of the net, and d is the distance through which the plankton net is towed.

De Bernardi (1984) [18] underlined the errors in the given calculation that can occur due to clogging of the net. Accordingly, use of nets with flow meters is strongly recommended, both for freshwater and estuarine mesocosms [6]. The best position for the flow meter was suggested to be in a position midway between the center and the net rim [27]. Moreover, a second flow meter outside the net can provide an estimate of net speed, and the two meters combined can yield an indication of filtration efficiency and clogging.

In case of using a flow meter, the volume of water that is sampled can be calculated using the specific formula, generally provided by the manufacturer of the flow meter [6].

7.4.3 Using water samplers to collect zooplankton in mesocosms

Several types of water or volume samplers (water bottles, tube samplers and pumps) can be used to collect (nano-micro- and) mesozooplankton while removing the zooplankton together with a known volume of water that will later be filtered to separate the mesozooplankton from the water sample. This approach has several advantages compared to net samples: 1) it does not change the relative composition of the organisms in the ecosystem by selective removal of organisms in the mesocosm, 2) the sampled volume can be precisely determined, and 3) for two of the water sampling types discussed below (water bottles and pumps) samples can be taken at higher spatial resolution than with most nets.

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Water bottles (Figure 7-2) sample relatively small amounts of water. Water bottles are thus generally “efficient for small, less motile organisms” which can “yield low collection efficiency when used to sample the larger, scarcer, and more active zooplankters” [18]. Thus, as relatively small amount of water is sampled, the number of replicates generally needs to be higher, especially to account for spatial heterogeneity of mesozooplankton distribution horizontally and vertically [18]. However, an alternative way to minimize spatial heterogeneity in mesocosms, is to mix the mesocosms thoroughly before sampling, or pooling samples from several locations in the mesocosms. Note that analysis time and cost can be reduced if single samples are mixed [1], rather than counted separately.

Figure 7-2. Examples of volume samplers that can be used for water and (zoo)plankton sampling: a) Ruttner bottle, b) Friedinger bottle [1], c) Niskin water samplers, and d) Van Dorn samplers [28], [29]. All have closure

mechanisms that allows sampling at desired depths [6].

Tube samplers can be seen as a “long water bottle” used to collect an integrated sample when lowered through the water column [18]. Using (rigid) tube samplers, the entire water column can be sampled (i.e. from the surface to e.g. a few cm above the sediment) with an appropriate diameter in shallow water mesocosms ([30], [31]). They include models such as Heart Valve and Limnos Samplers, described in Knoechel and colleagues (1992) and Pennak (1962) ([32], [33]). When using rigid tube samplers, one should consider that the length of the tube allows for sampling the wanted depth without being to long for easy handling [18]. It is also recommended in [1] to choose a sampler that is transparent (e.g. Plexiglas) to have a better idea for what and how much water is being sampled, allows the free flow of water when lowered and has a rapid and tight closing system.

When sampling, the tube should be lowered vertically to the predetermined depth at an even and moderate speed to eliminate most of the pressure wave in front of the opening [1]. The tube is then retrieved at a moderate speed. If the volume samplers are not used with an integrated net (ex. Bottles), the water should be filtered through a plankton net (or other straining equipment).

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To overcome heterogeneity in the mesocosms, it is advisable to pool samples from several locations in the mesocosm. The (sub)sample(s) should be filtered through a net with an appropriate mesh size, making sure that all material on the inside of the net is rinsed into the draining cup. The material should be transferred to the sample bottles and preserved or analysed as soon as possible after collecting on the net.

Excess water could be returned to the mesocosm if volumes are limiting, but returning water to the mesocosm that is sampled by nets may also introduce bias in organismal composition, that should also be considered.

Plankton pumps (hand pumps and motorized pumps), described e.g. by Nayar et al (2002) and Waite & O’Grady (1980) ([34], [35]) offers an alternative method to sample either at specific depths or over depth intervals in mesocosms. Sampling procedures and how to effectively use pumps to collect mesozooplankton were also summarized in De Bernardi (1984) [18], as quoted from Tonolli (1971) [26]. Compared to other volumetric samplers, pumps “permit the collection of the largest volumes of water” [18]. Accordingly, pumps can be employed as an alternative to bottles and tubes with some advantage if large water volumes are needed. However, “large active plankton species are liable to be sampled less efficiently using a plankton pump than by other types of quantitative samplers. The opposite can be the case for small species” [1], [18]. Use of “a large plastic funnel (diameter about 50 cm) at the end of the sampling tube” is recommended “to prevent escape of jumping copepods” [1]. Motorized plankton pumps with continuous flow-through are recommended rather than hand-powered plankton pumps, because motorized pumps provide a regular flow, thus providing better estimates of the quantity and composition of the plankton [1].

The approach used for quantitative estimation and small-scale distribution of plankton with pumps have been summarized in Kaisary and colleagues (2012) as: “an open-ended inlet hosepipe is lowered into the water and the outlet pipe is connected to a net of suitable mesh size. The net is particularly submerged in a tank of a known volume. This prevents damage to the organisms. The mesozooplankton is filtered through the net. A meter scale on the pump records the volume of water filtered” [6].

Sampling by gravity, can be used to collect mesozooplankton. Some land based mesocosm facilities allow taking water samples from a port or by a hose through gravity. In this case a sufficient amount of water needs to be collected (depending on density of organisms). The water sample is screened by an appropriate mesh (50-100µm) in the lab. Given that mesozooplankton is not retained quantitatively in the mesh, the screened water can be used for other analyses, e.g. chlorophyll-a and nutrient analyses, or may be added back to the mesocosm, if needed.

7.4.4 Additional materials generally needed for zooplankton sampling

Weight (for net sampling only, non-toxic materials should be considered in mesocosms). Squirt/Wash bottle to rinse out the net and the draining cup. Small plastic funnel for transferring the water sampled to sample bottles. Plastic (bucket) for pooling multiple water samples (e.g. from volume sampler). Filtration equipment to collect or concentrate samples from volume samplers. The filtration equipment may be either a plankton net, with an appropriate size to filter the targeted organisms or “a large funnel with draining cup fitted with a netting” [1]. Gloves (rubber or vinyl), waterproof notebook, waterproof pen, if ethanol or other solvents is used, an alcohol-proof pen or pencil is recommended for both internal and external marking, pipettes (to add preservative)

7.4.5 Sampling Containers and Sample Bottles

Bottles (100 ml, 200 ml or 250 ml), recommended to be (brown) bottles with screw-tops or glass vials for storing samples [1]. Select the right type of bottle for samples based on the type of the reagent used. For instance,

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plastics are not suitable for storing samples if Lugol’s Iodine is used as preserving solution [1]. See Appendix A for more details.

Labels, recommended to be from waterproof paper. As a best practice advice, mesocosm name, (treatments) date and time of sampling, the sampling depth, the sampling equipment used should be stated on the labels of each mesozooplankton sample [2]. Volume of water filtered, and the mesh size of the net used in filtering should also be recorded on the labels.

7.5 Sampling Design for Shallow and Deep Mesocosms

As a best practice advice, if the study aims to estimate species composition and abundance in the mesocosm, samples should be taken both from different depths of whole mesocosms; as the vertical and horizontal distribution of mesozooplankton is uneven.

Moreover, “if the objectives of the study require information to be collected regarding spatial variation in general, or a high level of accuracy in the estimates, it will be necessary to draw up a sampling design (programme) that is adapted to the” width and the depth conditions of the mesocosm [1].

For instance; “stratified, random sampling (see [36] for more information) of mesozooplankton is based on dividing the mesocosm horizontally and vertically into a series of sampling units (strata), from which a random selection of units to be sampled is selected. The optimum number of samples per stratum will depend upon three factors: the size of the stratum, the variability within the stratum and the cost of sampling in the stratum [1] ”.

In general, sampling should be carried out at an adequate distance from mesocosm walls and one shall be careful not to disturb the plants, if available.

7.5.1 Sampling time and frequency

Sampling time and frequency should be determined according to the main purpose of the study. There are many recommendations about time and frequency, such as

According to EN 15110 (2006), mesozooplankton should normally be sampled between 10 a.m. and 4 p.m. For studies of vertical migration, sampling every 6 hours is recommended, but as a minimum, sampling should be performed at midday and midnight, or at least roughly the same time each time to maintain consistency and allow comparison [1] .

“Many common species of macromesozooplankton (such as Chaoborus, Mysis) may only occur in the water column at night. If collection of these species is desirable, it is essential that sampling occur at least one hour after sunset” but it is best at midnight [2].

7.5.2 Shallow mesocosms

If mesocosms are mixed continuously, one composite (pooled) water sample (i.e., samples from different locations and depths can be combined) can be considered as representative for the entire mesocosm. If it is unclear whether the whole water volume is efficiently mixed, make sure to include all (thermally or salinity-based) stratified zones be covered.

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For mesocosms containing sediment and possibly with macrophytes, a detailed sampling design for collecting samples at multiple horizontal and vertical positions might be needed, as “large-bodied mesozooplankton (typically cladocerans) often aggregate within submerged plant stands or sediment during daytime (Timms & Moss (1984) [37]; Lauridsen & Lodge, 1996 [38], Tavşanoğlu et al. (2012), [39]), as macrophytes might serve as a refuge for mesozooplankton. In such mesocosms, mesozooplankton samples should be taken from both areas containing aquatic vegetation and from areas with little vegetation, just above the sediment surface without disturbing [1]. A suggested approach is summarized in [31] as follows:

“The entire water column, from the surface to approximately 5 cm above the sediment, is sampled from three positions in each enclosure: 10, 30 and 60 cm from the enclosure wall. Two samples are prepared: one to be used for chemical and phytoplankton analyses where water is sampled without touching the plants – and one to be used for mesozooplankton analyses, where water is sampled also close to the plants”.

As sampling device for shallow and macrophyte-covered mesocosms, use of a volume sampler, plankton pump or flexible tube (i.e. core/tube sampler) with an adequate diameter (i.e. > 6 cm) to sample from the entire water column are recommended [1]. If tube sampler is not available, samples can be taken with “Ruttner water sampler from the surface (i.e. 20 cm below the water surface), middle and the bottom (i.e. 20 cm above the sediment). The sample in the middle should be adjusted according to the enclosure type and actual water depth” [31]. In addition, a heart valve or Limnos sampler can be used for sampling.

7.5.3 Deep mesocosms

If the water column in the mesocosm is stratified or partially mixed, the sampling procedure must be determined after careful considerations on the water layer(s) to be sampled (e.g. sampling discrete water layers; sampling multiple depths with subsequent pooling to one combined representative of a layer).

✓ As best practice advice, a pilot study prior to actual study can be carried out to examine mesozooplankton densities at several horizontal and vertical sites in a mesocosm (two or three locations previously determined, in line with the purpose of the study. The results of the pilot study can be used in planning the actual sampling design.

✓ Sampling design and the estimation of mesozooplankton sampling precision are reviewed by Green (1979) [40], Downing et al. (1987) [41], Eberhardt and Thomas (1991) [42], Pace et al. (1991) [43], and Prepas (1984) [44] in more details.

✓ Absence of vertical temperature profiles indicates vertical mixing, but NOT necessarily homogenous distribution of motile organisms like zooplankton. Heterogeneous distribution of mesozooplankton should be assumed as the mesozooplankton densities and species composition vary vertically and horizontally in water bodies and has not been proven to be (practically) homogenous.

✓ Diel vertical migration (DVM) has been shown to be an important anti-predator defense mechanism in deep stratified mesocosms (in Gliwicz (1986) [45] and Lampert (1989) [46]) that can be taken into account for sampling.

According to the European standard EN 15110 (2006), “samples should be taken at intervals of no more than 2 m in the epilimnion and metalimnion” [1] . Accordingly, in deep mesocosms (i.e. 20 m depth), the samples taken should represent the mesocosm vertically and horizontally. The composited samples (i.e. integrated samples

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taken from a given range of depths) should include the epilimnion and the hypolimnion. An example methodology is summarized in Berger and colleagues (2010) [47], for sampling mesozooplankton from deep mesocosms. As a sampling device, for deep mesocosms, it may be more appropriate to use sampling equipment that allows efficient sampling of the whole water column, such as plankton net with a closure or a plankton pump rather than volume samplers, especially for practicality and safety issues [1] . If vertical net hauls are used, all depths should be covered. Remember that the net can be clogged easily in case the mesocosm includes massive quantities of algae and other particles. However, for sampling the entire water column, a flexible hose as tube sampler may be preferable to all alternative methods due to highest sampling efficiency (Walles and Nejstgaard, in prep).

7.5.4 Best Practice Advices on selecting the right sampling instrument

In order to provide a pathway to select the right sampling equipment for quantitative mesozooplankton sampling, De Bernardi (1984) compiled the following table from several comparison studies (Table 7-2). For further information on the selection of the appropriate instrument, Gehringer & Aron (1968) [27], Tonolli (1971) [26] and Bottrell et al. (1976) [48] should be visited [18].

Table 7-2. “Schematic recommendation for the choice of sampler to be used for the assessment of mesozooplankton population density under various conditions” (Redrawn from [18]).

Deep and pelagic Shallow and littoral

Point samples

Vertically integrated

samples

Point samples

Vertically integrated

samples

Samples within

vegetation

Ruttner, etc. + - + - +

Traps ++ - ++ - +

Tubes - ++ - ++ ++

Pumps ++ + ++ - ++

Nets - ++ - - -

Plankton samplersa

- ++ - - -

a i.e. Clarke-Bumpus

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7.6 Best practice advice on preservation and storage

Samples for quantitative analyses should be immediately preserved. If this is not possible,

1) non-preserved samples should be kept cold (2 - 5 °C) and dark as short as possible, as larger organisms may alter the content rapidly by artificial predation rates in the concentrated sample [1].

2) preserving fluid can be added to the sampling bottles in advance. If preserving fluid has not been added to the sampling bottles in advance, it should be added immediately after the samples have been taken.

3) To minimize shrinkage and muscular tension, zooplankton organisms may be anaesthetized before preservation, e.g. with carbon dioxide in mineral water.

Ethanol is a preferred fixative as it is less toxic than e.g. formaline and allows better molecular analysis of the samples. If ethanol is used, the preserved sample should have an ethanol content of at least 70 % to 75 %.

Samples for genetic analysis based on DNA extraction should be stored in a refrigerator or freezer, but in such a way that the sample itself does not freeze, which is ensured by preservation in high concentrations of ethanol [1]. For such samples, ethanol with a concentration > 90 % is recommended (an ethanol content higher than 90 % is recommended). An aqueous solution based on pure ethanol should be used for this purpose (instead of methylated spirit).

Note that samples for genetic analysis based on electrophoresis should be stored in a freezer (approximately -75 °C) and should not have any preserving solution added to them [1].

A high ethanol concentration helps to maintain body shape (Cladocerans) as well as preventing the animals from releasing their eggs.

If formaldehyde is used, the preserved sample should have a formalin concentration of around 4 %, i.e. 20 ml aqueous 20 % formalin should be added per 100 ml sample volume, or approximately 10% if concentrated formaline (37%) is used.

Preservation in a cold solution of formalin (6 °C) with added sucrose reduces the chances of the animals releasing their eggs.

If Lugol’s Iodine is used, add in the amount of 0.5 ml to 1.0 ml per 100 ml sample volume. While Lugol’s has the advantage of being non-toxic for humans, it is also a weak preservative. Thus, samples containing large quantities of animals and other organic material (particularly samples collected in littoral regions) require addition of more Lugol’s Iodine than samples that contain little organic material; e.g. 3 ml to 5 ml per 100 ml sample volume. Samples preserved with Lugol’s Iodine should always be stored in the dark and preferably chilled to below 5 °C, unless they are to be analysed within a week, in which case they can be stored in the dark at room temperature [1]. Samples that have been preserved with Lugol’s Iodine should be straw colored and should be checked after a couple of days for oxidation” [1]. The preservative should be renewed every 6 months for better storage.

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8. Quantitative Analysis of Zooplankton

8.1 Sample Preparation

8.1.1 Subsampling

Zooplankton samples frequently contain many organisms and it is often impractical to count every individual. A number of methods have been developed to subsample mesozooplankton collections.

An example subsampling process by pipettes was summarized in [49] as follows:

1. Use a swirling flask and Hensen-Stempel pipette (i.e. the pipette, which is a wide bore piston apparatus; ensuring the organisms not to be selected against based on size, that can capture a 2.5 to 50 ml volume of liquid). ✓ Edmondson and Winberg (1971) suggest that the opening of any pipette used to take subsamples,

whether automatic or manual, should exceed 4 mm [17]. ✓ Since the volume of the pipette is fixed, the volume of the sample should be altered to give

subsampling densities > 60 individuals per subsample. 2. The sample is accurately brought to the desired volume and then poured into the flask. 3. It is then swirled in a figure eight pattern until mixed. 4. While in motion, the Stempel pipette is inserted into the flask and the subsample is taken.

✓ In [48], it is found that the coefficient of variation for subsampling in this way stabilized at 0.08 when the density of organisms in the subsample exceeded 60 individuals.

Another subsampling process by Plunger sampler is as follows:

“The sample is filtered at an appropriate mesh size, which is smaller than the one used in the field (for instance,

20 µm). The content is rinsed into a sampler (as in Figure 9-1) and it is filled until 100 ml. The piston (Plunger) is put into the sampler (the tube). Make sure that the plunger sampler is placed vertically and straight in the sampler and the lid is tight. The water in the sampler is mixed carefully and gently by turning the sampler upside down a number of times. Place the sampler on the table and drag up (gently, but quickly) a known volume (i.e. 2.5-10 ml) of sample. Depending on the density of the animals, several subsamples, up to 40 ml in total, could be taken from one sample and added into one counting chamber. Pour the subsample in a counting chamber and rinse the piston into the counting chamber” (Personal communication with Liselotte Sander Johansson, AU, and from [50]).

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Figure 8-1. The Plunger Sampler (Adopted from [50])

A wide variety of plankton splitters have also been developed for the subsampling of mesozooplankton samples (e.g. Kott (1953) [51]), which are generally made of plastic with internal partitions. Folsom plankton splitters are widely used (in marine research) and by which, the mesozooplankton samples are divided into equal fractions. Kaisary and others (2012) summarized the subsampling methodology with a plankton splitter as follows:

“The mesozooplankton sample to be sub-sampled is poured into the drum and the drum is rotated slowly back and forth. Internal partitions divide the samples into equal fractions. The fraction should be poured again into the drum for further splitting. The process is repeated until a suitable sub-sample is obtained for counting. The splitter is thoroughly rinsed to recover the organisms, which should be sticking onto the wall of the drum. The sample is usually splitted into 4 sub-samples. One of the sub-samples is used for estimation of dry weight, the second for counting the specimens of common taxa, the third for relative abundance of species and the fourth fraction is kept as reference collection. Plastic or glass pipettes are also used to take the sub-sample for counting. The Stempel pipette is used to obtain a certain volume (0.1 to 10 ml). The mesozooplankton sample in a glass container is diluted to a known volume and is stirred gently. The Stampel pipette is then used to remove the sub-sample or aliquot for counting” [6].

8.1 Sample preparation for counting

Several methods can be used to remove organisms from a water sample, such as sedimentation, centrifugation, and filtration. According to De Bernardi (1984) [18], filtration is the most convenient way however, Bottrell et al. (1976) [48], suggest that sedimentation is the best method for rotifers, even though it is practical only at high densities.

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✓ One shall note that, with sedimentation it is always laborious and difficult to determine whether all the organisms have settled. The complete settling of suspended material can take several days. In addition, removal of the supernatant liquid by siphon requires much care to ensure that animals are not disturbed [18].

Cautions on filtering (adapted from [18]):

● The filtration must be done at low pressure and as gently as possible to avoid damage to the fragile organisms or their forced passage through the mesh.

● Rigid filters are preferable because their mesh size is invariable. ✓ Likens & Gilbert (1970) [52] suggest the use of nets with a mesh aperture of 35 µm which,

according to the authors, permits an appropriate retention of even the smaller rotifers. ✓ On the other hand, Schindler (1969) [53] and Ejsmont-Karabin (1978) [54] indicate that nets

with 28 or 10 µm mesh sizes are the most suitable for collecting the smallest forms especially from nutrient poor mesocosms.

8.2 Counting Procedure (Enumeration)

8.2.1 Counting chambers

Rotifers and nauplii are counted using either a Sedgewick-Rafter cell or a sedimentation chamber (1-25 ml) [17]. Sedimentation chambers can be preferred since these allow more flexible volume changes than Sedgewick-Rafter cells, whereas Makarewicz & Likens (1979) [55] found no difference between counts obtained using these two chambers.

In addition, rotifers and nauplii can be enumerated in a 1 – 5 ml clear acrylic plastic counting cell fitted with a glass coverslip [6].

Crustaceans are usually counted using a counting chamber (5 – 10 ml) “with partitions or grooves which confine the subsample to tracks of a constant width” [49]. The width of the grooves in the tray should be less than the width of the field at the counting magnification. Petri dish with grid lines on the bottom, a grind edge and a glass cover to avoid air bubbles, preventing the animals from moving is also usable. Bogorov design-based counting chambers for mesozooplankton can also be used for counting. An example counting chamber can be examined in Hydrobios website (REF: https://www.hydrobios.de/produkt/zahlkammer-fur-mesozooplankton/?lang=de).

A Sedgwick-Rafter cell is not suitable because of size of the crustacean [6]. According to Kaisary and others (2012), an open counting chamber “80 by 50 mm and 2 mm deep is desirable; however, an open chamber is difficult to move without jarring and disrupting the count” [6].

✓ As a best practice advice; if the counting chamber do not have partitions or grooves, mild detergent solution can be placed on the chamber before counting to reduce the movement of organism

It should be noted that, the chamber for counting the sample may vary with the type of microscope used.

8.2.2 Counting (enumeration) methodology

✓ “Samples should contain a minimum of 200 animals (for crustaceans: exclusive nauplii) to provide a good estimate of numbers and species composition. If both crustaceans and rotifers are included in the analysis, the samples should contain a minimum of 200 animals of the group, which has the fewest individuals” [1] .

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✓ If samples from different dates are pooled, it would be a good idea to use half of the sample for pooling and keep the rest as a date specific sample.

McCauley explained how to count the rotifer and nauplii as follows: The rotifers and nauplii

“are allowed to settle, approximately 2h per cm height. After settling, the chamber is placed on the stage of the inverted microscope (i.e. Ütermohl technique) and individuals counted. It is best that the contents of the whole chamber be enumerated to simplify statistical considerations. This can be achieved by altering the volume of liquid in the sample and the subsample to bring the number of organisms in the chamber to the desired level. It is important that all manipulations of sample volume are recorded and are as accurate as possible so that the original concentration of organisms in the sample may be calculated” [49].

Counting crustaceans,

“Once the subsample is placed in the chamber, the individuals are counted by moving the chamber and tallying the individuals encountered in the field of the microscope. Since all of the organisms in the subsample are usually counted, a homogeneous distribution in the chamber is not needed, although the volume of the sample should be adjusted so that organisms do not pile up on one another. Repeated subsamples are then taken until the desired number of individuals have been counted” [49].

✓ An excel macro including the sheets that can be used for counting individuals of different species and to make species-area curves (saturation curves) of counted individuals to species is provided as an attachment to this document (EXCEL 2 – Counting mesozooplankton.xmls). Information on how to use the excel macro is provided in Appendix C.

✓ A high-quality (binocular) dissecting microscope is sufficient for counting Crustacea, although a light microscope is required for taxonomic identifications [6].

Zooplankton are enumerated in gridded chambers and Bogorov chambers that prevent duplication of counts. The number of animals counted will vary with the desired degree of sampling precision and the goals of the study [44].

An example counting (enumeration) procedure was summarized in [31] as follows:

“mesozooplankton is counted using a stereomicroscope until at least 50 individuals of the most dominant species have been counted [31].

Another counting (enumeration) procedure was summarized in [6] as follows:

“It is recommended that the sub-sample or an aliquot is taken for the common taxa. For enumeration of mesozooplankton the sub-sample or aliquot of 10 to 25% is usually examined. However, the percentage of aliquot can be increased or decreased depending on the abundance of mesozooplankton in the sample” [6].

✓ “For the rare groups, the total counts of the specimens in the samples should be made” [6].

8.3 Best Practice Advice on subsampling and counting

✓ Before subsampling, all large organisms such as fish larvae, coelenterates, decapods and other should be removed and enumerated [6].

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✓ It is essential to determine whether subsampling conforms to a random distribution statistically, since this information is required in determining estimates of counting precision. To confirm that subsampling is random, replicate counts should be compared with a Poisson distribution using a Chi-square test ([2] and [49]).

o Lund et al. (1958) [56] recommended that at least 10 sets, each containing > 5 replicate counts, should be tested for randomness. It is important to test subsampling procedures on a variety of samples if the density of individual organisms varies considerably among the samples to be enumerated. If the counts of discrete plankton organisms from a sample are random and the number of individuals counted is small relative to the total population then, the counts can be assumed to be distributed according to a Poisson series. If the counts are not random, then the entire sample should be counted [49]. The entire samples should also be counted without subsampling in case the mesozooplankton density is low (<200 zooplankter) [6].

✓ The coefficient of variation stabilization technique, that should be utilized to determine population abundance of a number of taxa simultaneously was summarized in [6] as follows:

“a given sample is split into sub-samples until the 3 most numerous species are present in numbers of at least 30 animals per sub-sample. All organisms in the sub-sample are then counted, including all rare species. Alden et al. (1982) supply a table in which the count for each species in this sub-sample can be looked up, and the sub-sample needed to obtain a count of at least 30 individuals per species can be read off the chart (i.e., the ½ sub-sample, the ¼ sub-sample, 1/8, 1/16, 1/32, etc.). Thus, each of the rare species can be counted to the minimum required precision (i.e. 30 animals per sub-sample) by counting only one more sub-sample. Counting 30 animals per sub-sample gives a 95% confidence limit of ±30% of the mean” [6].

● With this technique, the abundance of common species to a predetermined level of precision can be estimated ([2] and [49]).

● “The coefficient of variation is equal to the inverse of the square root of the number of

individuals counted (CV = 1/√𝑁). For example, if 50 individuals were counted then the coefficient of variation would be approximately 0.14” [49].

● “This relationship can be used to determine how many individuals should be counted to obtain a desired level of precision for the estimate of the density of organisms in a sample” [49].

● Table of calculated values of the coefficient of variation (expressed in fractional form) of biomass estimates can be used to estimate the number of individuals that should be counted and to indicate when increases in counting effort are advantageous. For the table, please visit [49]. For example, moving down a column at a fixed coefficient of variation indicates the gain in precision with successive increases of counting effort.

✓ It should be noted that, the calculated values of precision presented in the mentioned table are based on a number of assumptions and should not be used without determining whether or not these assumptions are true for processing a particular sample.

8.4 Taxonomy and nomenclature

The list of important studies in identifying mesozooplankton are provided in Table C-1 at Appendix D.

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9. Estimating the biomass

✓ larger zooplankters (such as medusae, ctenophores, salps, siphonophores and fish larvae) should be removed from the mesozooplankton sample and their biomass should be estimated separately. The total biomass, then, will be the biomass of larger plus the biomass of the rest of the mesozooplankton [3].

✓ To estimate mesozooplankton biomass, length measurements should be made for minimum 30 rotifers, 30 cladocerans, 30 copepodites and 15 adult copepods per species [31].

9.1 Crustacean Zooplankton

9.1.1 Measuring the length or dimensions of an organism

Length (or dimensions) of a crustacean mesozooplankton can be measured using an ocular micrometer (a dissecting microscope equipped with an ocular micrometer) or a computerized measuring system [49].

✓ It should be noted that the use of an ocular micrometer reduces, but does not remove, the possibility of error in measurements due to variation in the angle and distance between the eye of the observer and the eyepiece of the microscope [49].

✓ Length of some of the crustacean mesozooplankton is dependent on its developmental stage. ✓ In addition, with the help of digitized plankton images, the analysis can be done using the commonly used

image analysis softwares, such as Zoo/PhytoImage software (licensed under GNU/GPL) (ZooIMAGE) [57] and ZooScan [58].

For more information on this technique, please refer to [59]–[61].

It should be noted that definition of total length can vary according to the type of the mesozooplankton. Below are some examples on how to measure the total length:

● The total length of Cladocera can be measured from the top of the head to the point of insertion of the tail spine. Measurement starting from the eye region to the point of insertion of tail spine is also used commonly among length measurements of Daphnia [49].

● In case of Copepoda measurements, the length can be measured from “the tip of the cephalothorax to various points on the urosome, usually excluding the furcal rami” [49].

● McCauley (1984) [49] also noted that, in measuring the length of marine Copepods, maximum width of the body can be used rather than the length in order to avoid ambiguity in measurements.

✓ For an analysis of the precision of estimates (errors in measurements), please refer to [49].

9.1.2 Length-weight relationships (Adopted from Bottrell et al. (1976) [48])

Paterson (2001) claims that the most common method for estimating the dry weight of mesozooplanktonic Crustacea “relies on converting estimates of length to biomass using length-weight regressions” [2].

✓ As best practice advice, results from measuring systems can be combined with an electronic database to ease data entry and storage (e.g. Allen et al. (1994) [62]).

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An excel sheet, including the length-weight relationships for crustacean mesozooplankton will be provided as an attachment to this SOP (Excel_1, in preparation). (See Appendix E for the list of references that will be cited).

✓ Length of copepods can be converted into biomass using conversion factors by Paffenhöfer and Harris (1976) [63], Landry (1983) [64] and Castellani et al. (2005) [65].

9.1.3 Estimating the dry weight (Adopted from [49])

1. “To determine the dry weight of crustaceans, individuals are isolated from the sample using a pipette or a fine dissecting needle (See Furnass & Findley (1975) [66], that presents an apparatus to sort individuals without picking them up manually, which can be attached to the stage of the dissecting microscope)

✓ If the weights are for length-weight relationships, then eggs embryos, and gelatinous sheaths should be removed prior to weighing ([67], [68]).

2. The decision on whether to weigh individuals or groups of animals can be made based on the expected weight of an individual.

✓ Dumont et al. (1975) [67] suggested that the minimum amount of weighed material should exceed 5 µg, given that the sensitivity of the most commonly used balances.

3. The animals should be rinsed with distilled water if they are from preserved samples. The animals are then placed on tared aluminium foil pans or boats ([67], [55]).

✓ The material used as a weighing pan varies among studies. Whatever the material used, the weight of the pan should be kept to a minimum ([17]).

4. The animals, on their trays, are then placed in an oven and dried for 24-48 h (varied between 2 and 48 h with 24 h being the most frequently used) at 60°C (as in [55], [68]–[72]).

5. After drying, the samples are allowed to cool in a desiccator for 1-2 h and are then transferred to the balance and weighed using counterbalance techniques to maximize the accuracy of the reading. A small container of desiccant should be placed in the chamber of the balance to absorb moisture picked up while transferring the sample.

✓ Placing a container of desiccant inside the housing of the balance and sealing this space also improves the stability of the reading.

✓ Weighing should be delayed for a short period of time, to allow for stabilization.

To obtain an estimate of precision, replicates of individuals or groups can be weighed. The number of replicates can then be varied to give a desired level of precision. Please see [73]–[75] for more details.

9.2 Rotifers

9.2.1 Predicting dry weight using geometrical formulae (Adopted from [49])

For Rotifers, geometrical formulae are usually combined with appropriate length measures to estimate biomass.

Estimating the volume of Rotifers, “several different dimensions are measured and then used to calculate a volume estimate from simple formulae which depend upon the shape of the species (i.e. geometrical formulae)”.

✓ The geometric formulae provide accurate descriptions of the volume of individuals [49]. ✓ A complete description of the technique as well as the list of geometric formulae to calculate the volumes

of over 20 genera can be found in Ruttner-Kolisko (1977) [76] and McCauley (1984) [49] (can be provided in Excel sheet, Excel_2).

The length of rotifers can be measured using inverted microscope.

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✓ The dimensions needed to be used in geometric formulae should be estimated on “slightly narcotized, living adult specimens, that have not been flattened by a coverslip” [49].

Once the volume has been calculated, it is converted to fresh weight assuming a specific gravity of 1, as the specific gravity of all species in all habitats equals one [49].

Fresh weights for various Rotifers, which were determined based on the geometric volume calculations, are presented in Bottrell et al. (1976) [48]. Given fresh weights can be converted to dry weight, as the dry weight is a constant fraction of fresh weight irrespective of the species being considered, by assuming some constant value [49].

✓ Constant value was assumed as 0.05 (dry:wet) in [77] and 0.1 (dry:wet) in [70] and [78]. Measuring the dry weight by microbalance or weighing is impractical and inefficient for rotifers. As the small size of rotifers makes it impossible to weigh individual animals of most species [49]. They should be weighed as a group and a mean weight should be determined by accounting for the number of individuals used [49].

✓ Ruttner – Kolisko (1977) [76] reminded that this technique requires more time and less efficient in determining weight than volume and weight determination with geometric formulae.

If considered, [55] and [67] described this procedure clearly. It is also summarized by [49].

9.3 Estimating Mass as Carbon Content (Adopted from [49])

The technique proposed by Latja & Salonen (1978) [79], to determine the carbon content of individuals “requires high temperature combustion of the sample and measurement of the carbon dioxide evolved using an infra-red gas analyzer. It requires 10000 times less material and processes samples 20 times faster than current methods of calorimetry” [49].

“Using this technique, sub-microgram quantities of carbon can be measured. This sensitivity is considerably better than the best microbalance, and the carbon content of an individual animal as small as a rotifer can be determined in < 1 minute” (as in [79] and [49]).

The technique was summarized by [49] as follows:

1. “An individual is transferred to the combustion tube using forceps, and then combusted at 950°C. ✓ If the animal is transferred with a pipette, then the carbon content of the volume of water used

must be determined and subtracted from the weight of the animal. 2. The carbon dioxide produced is measured using an infra-red gas analyzer. A complete description of the

apparatus, its calibration, recovery statistics, and design for constructing the quartz combustion tube can be found in Salonen (1979) [80]” [49].

3. The probable conversion factor (µg C per µg dry weight) for different taxa can also be used and can be reviewed from [81] and [82].

9.4 Estimating biomass: other methods in literature

The biomass can also be estimated by the following methods (as provided in [6]):

● Volumetric (displacement volume and settling volume) method (in the field or laboratory): The total mesozooplankton volume is determined by the displacement volume method. In this method; a) “the mesozooplankton sample is filtered through a piece of clean, dried netting material. b) the mesh size of netting material should be the same or smaller than the mesh size of the net used

for collecting the samples. c) The interstitial water between the organisms is removed with the blotting paper.

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✓ While blotting, due care should be taken not to exert too much pressure as to damage the delicate organisms or specimens.

d) The filtered mesozooplankton is then transferred with a spatula to a measuring cylinder with a known volume of 4 % buffered formalin.

e) The displacement volume is obtained by recording the volume of fixative in the measuring jar displaced by the mesozooplankton.

f) The settled volume is obtained by making the sample to a known volume in the measuring jar” [6]. ✓ Using this technique, the plankton is allowed to settle for at least 24 hours before

recording the settled volume [6]. ● Gravimetric (wet weight, dry weight and ash free dry weight) method (preferably in laboratory): In

this method: a) “Filter the mesozooplankton and remove the interstitial water by a blotting paper.

✓ While blotting, due care should be taken not to exert too much pressure as to damage the delicate organisms or specimens.

b) The mesozooplankton weight is taken on predetermined or weighed filter paper or aluminium foil. Express the net weight in grams”. c) The dry weight method is dependable as the values indicate the organic content of the plankton.

Analysis such as the dry weight is determined by drying an aliquot of the mesozooplankton sample in an electric oven at a constant temperature of 60ºC.

✓ The whole or total sample should not be dried because subsequent analyses such as enumeration of common taxa and identification of the species wouldn’t be possible after drying the sample.

d) The dried aliquot is kept in a desiccator until weighing. Express the values in milligram. e) Ash free dry weight method is also occasionally used for biomass estimation” [6].

✓ This method can be used for estimating the biomass of gelatinous mesozooplankton.

10. Size Distribution

Differences in mesozooplankton size distribution can be assessed by calculating size diversity and normalised biomass-size spectrum (NSS).

10.1 Size Diversity

Size diversity can be calculated by using “function of probability density of individuals with respect to size” [83] as described in [84], [85]. As it is a continuous function, this approach provides the advantage of avoiding “the arbitrariness introduced when using size classes” [83].

Moreover, Quintana and colleagues (2008) provided the methodology, which is based on the Shannon-Wiener diversity expression [86] and adapted to a continuous variable, such as the body size; to estimate the size diversity by using the individual size measurements [87].

● One should note that, if size diversity is high, whether the size range (i.e. the range representing the minimum and maximum length of species in µm) is wider and/or there are similar proportions of the different sizes [83], [87]–[89].

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10.2 Normalised biomass-size spectrum (NSS)

As stated by Brucet and colleagues (2010), “biomass-size spectrum describes how biomass of organisms is distributed along size classes” [90]. One should see the book by Kerr and Dickie, published in 2001, ‘The Biomass Spectrum: A Predator-Prey Theory of Aquatic Production’ [91] for more details on how to assess the size distribution with NSS approach.

As a best practice advise, using size diversity approach is advantageous over NSS as the former one does not require statistical analysis (in contrast to NSS) and “is represented by a single value” [92].

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11. Quality assurance and quality control

According to [93], systematic or random errors can occur during the sampling operations. Systematic errors are generally due to the “poor sampling practices or equipment design failures” which are usually constant [93]. On the other hand, random errors are generally unavoidable or unpredictable. One shall follow an effective Quality Assurance/Quality Control (QA/QC) strategy during the experiment to identify, quantify and control the errors. Standardization of sampling and analysis methods, taking replicate samples and analyses, or following a laboratory accreditation scheme are some examples for QA practices that can be followed during the experiment. Quality Assurance (QA) strategies are defined by the scientist-in-charge to ensure that the sample data meets Data Quality Objectives (DQO), which shall be defined prior to sampling. On the other hand, Quality Control (QC) is “the system of guidelines, procedures and practices designed to regulate and control the quality of products and services, ensuring they meet pre-established performance criteria and standards”. The QC practices that can be followed are as follows: taking “sample blanks, replicates, splits”, having and following “equipment calibration standards”, determination of “sample container size, quality, use and preservative amount” prior to sampling. The WISER project results highlighted the information below for QA of sampling and counting of plankton:

“1) Details of microscopes, chambers (individually identified and calibrated) and calibration of all ocular/objective combinations should be recorded in a note book and kept for reference. If fixed volume pipettes are used, these should be calibrated annually. 2) Checks for random distribution of sample should be done visually at low magnification for each sample. Some simple checks include: Comparing the number of observations in:

(a) half a chamber with the other half (b) comparing counts in the 1st transect with the 2nd transect (c) comparing counts in the first 20 field of view with the next 20 fields.

A more detailed check using simple Chi squared test should be done if a sample does not appear to be randomly sedimented or 1 sample every 3 months or so” [94].

According to the ‘Common Implementation Strategy for the Water Framework Directive (2000/60/EC)’, the intercalibration exercises in between laboratories will provide a “continuous quality assurance system”, by ensuring the results meet targeted levels [95]. In order to perform intercalibration in the AQUACOSM community, there shall be QA measures in each of the mesocosm facility. The common QA measures, that are determined based on this SOP and the valid sampling and analysis methods used during the experiments, that shall be taken by each mesocosm facility are listed as follows:

● “Field sampling and sample label

● Sample storage and preservation;

● Laboratory analysis” [95]. Further suggestions to assure sampling quality (not specific to mesozooplankton analysis) are as follows:

● One shall establish of a “routine internal quality control” and participate in “external quality assurance (QA) schemes” [95].

● In addition, one shall consider the resolution/degree of identification in counting and take pictures of unidentified species or species which have unsure identification.

For detailed QA methodology provided for sampling, analysis and counting of mesozooplankton, please visit EN 14996:2006 [96] and EN 15110:2006 [1] .

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12. Appendix

Appendix A

Table A – 1. The advantages and disadvantages of preservatives (Tabularized from [1] )

The preservative solution

Advantages Disadvantages

Lugol’s Iodine ● It enters the cell more quickly than formaldehyde, leaving shock-sensitive organisms better preserved in the sample.

● Detection of small mesozooplankton is easier, thanks to the enhanced contrast between organisms and the surrounding fluid.

● Slightly toxic in normal conditions of use and offers minimal discomfort to laboratory personnel.

● The intense staining may obscure certain cellular structures that need to be observed for proper identification (e.g. surface structures). Overstaining can be cleared up by adding sodium thiosulphate, which reduces the iodine.

● Organic matter is not fixed, and soft materials may lose their characteristic structure during storage of the sample. This alteration can be prevented by the addition of formaldehyde.

● Iodine is oxidised over a period of time; therefore, when storage time is long or samples contain a large amount of organic matter, samples need attention to prevent them from decay.

Formaldehyde (Formalin)

● Effective prevention of the microbial degradation of organic matter.

● Organic structures and other morphological characteristics remain visible.

● When stored properly in suitable bottles, samples will stay in good condition for many years without attention.

● Distortion of the body structure takes place especially in soft-bodied organisms (e.g. several rotifers and cladocerans)

● Cladocerans (e.g. Daphnia, Bosmina. Diaphanosoma) may balloon followed by the loss of brood-pouch contents (eggs and embryos). This can be prevented by using a cold (6 °C) solution of formalin with added sucrose.

o In this case, it is advisable to kill the organisms using more rapid and efficient methods or to utilize special solutions

and techniques [18].

● Formaldehyde is irritating at even very low concentrations. Formaldehyde can also be carcinogenic. This preserving solution should therefore be handled with care, and should be “washed out” of samples within an air extraction hood before they are analysed.

Ethanol ● The quality of the samples can be retained for long periods of time if they have been stored correctly.

● Use of 96% ethanol prevents carapace distortion and loss of eggs and embryos due to ballooning in cladocerans

● DNA is retained in a form which can subsequently be extracted for genetic analysis.

● Cell shrinkage can be observed, which will result in underestimation of dimensions.

● May be unpleasant for laboratory personnel (dizziness, headaches). For this reason, this preserving agent should be diluted before samples are analysed.

How to prepare the reagents?

a) Lugol’s solution (Adopted from [1] )

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✓ Distilled/demineralized water

✓ Potassium iodide (KI)

✓ Iodine (crystalline)

✓ Glacial acetic acid

Lugol’s Solution preparation procedure

Dissolve 100 g KI (potassium iodide) in 1 l of distilled or demineralized water; then add 50 g iodine (crystalline), shake until it is dissolved and add 100 ml of glacial acetic acid. As this solution is close to saturation, any precipitate should be removed by decanting the solution before use.

b) Formaldehyde Solution Preparation Procedure (Adopted from [1] )

Formaldehyde is neutralized, e.g. with hexamethylenetetramine (C6H12N4). Dilute the formaldehyde with water to 20 % (v/v) to avoid precipitation, and then add 100 g of hexamethylenetetramine and 40 g to 80 g sucrose per litre of 20 % formaldehyde.

c) Buffered sucrose formation (Buffered Formalin) (Adopted from [1] and [5]):

✓ Sucrose (crystalline),

✓ Formalin (37% solution of formaldehyde in water),

✓ Borax (powder)

Buffered Formalin preparation procedure

Dissolve 60 grams of sucrose in 1000 mL of formalin [97]. Then, dissolve 9 grams of borax in 1000 mL of sucrose formalin. Store in a labeled plastic container.

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Appendix B

Appendix C

Counting cladocerans

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✓ Open the excel file “Counting_cladocerans” on the computer and make sure the macro is enabled.

✓ Take some time to get acquainted with the file.

Note: On “Sheet 1” there’s a row with 50 cladoceran species, a column for the input of sample codes and 14 buttons. These buttons are all working as soon as the macro is enabled, but to activate or deactivate a quick key function of the buttons with the numbers 1-9 and “num enter” you can use Control+Q or Control+W respectively, or you can click on the buttons “Activate Quick Keys” or “Deactivate Quick Keys”. Once activated the buttons 1-9 correspond to the keys 1-9 on the keyboard and the numpad of the computer and the button “num enter” corresponds to the enter key on the numpad. The buttons can be dragged and dropped to a different location by right clicking to select them and then left clicking to drag and drop them.

✓ Pressing a button will enter +1 to the cell corresponding to that button and will also enter data in “Sheet 2” (more information on “Sheet 2” follows). Which cell corresponds to the button depends on the location of the button and the location of a preselected cell. The column the button resides in is the column the data will end up in and the preselected cell determines the row of the cell the data will end up in (this can be any cell in the row).

✓ Each time a button is pressed in “Sheet 1” “Sheet 2” counts the number of species and individuals in the selected row and stores that information in two columns, aptly named “COUNT_SPECIES” and “SUM_INDIVIDUALS”. These two data points are then added to a graph right next to the two aforementioned columns, thereby creating a saturation curve.

✓ “Sheet 3” named “ml” has columns for e.g.: recording the starting volume of and the amount of the subsample taken from the sample and “Sheet 4” named “All_Saturation_Curves” is there for transferring the two columns of “Sheet 2” to, after a sample is finished.

✓ Activate the quick keys and start counting the cladocerans. Drag and drop the buttons 1-9 and “num

enter” to the columns of the most encountered species for optimal speed.

✓ Use the two dissection needles to turn and if necessary to dissect the cladocerans.

✓ When the identification of a certain individual with the stereomicroscope is impossible, use the glass

Pasteur pipette to transfer the individual to the microscope depression slide. Add a drop of water to fill

the depression and use the tissue to remove access water after covering the depression with the cover

slip.

✓ After identification transfer the individual from the slide to the small third Petri dish.

✓ When all the individuals in the subsample have been counted, transfer the contents of the gridded (and

the small, if the subsample counted was the last one) Petri dish to a clean culture tube labeled

“sample_name done”.

Note: check with the stereomicroscope if all mesozooplankton was transferred, often some get stuck to the bottom of the Petri dish.

✓ Continue taking subsamples and counting until the saturation curve (see “Sheet 2” of the excel file

“Counting_cladocerans”) levels out. Since leveling out is a somewhat flexible concept, counting 100

individuals without finding a new species is hereby set as the definition of this concept. This means

counting can be stopped as soon as a whole subsample has been counted and no new species were found

for the last 100 individuals.

✓ When starting a new sample make sure that all materials are clean of mesozooplankton by thoroughly

rinsing them.

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Appendix D

Key literature to identify Crustacea:

● L. A. Błędzki and J. I. Rybak, “Freshwater crustacean mesozooplankton of Europe,” Switz. Springer, 2016. [98]

● L. M. Witty, “Practical guide to identifying freshwater crustacean mesozooplankton.” Cooperative Freshwater Ecology Unit, 2004. ([99])

● U. Einsle, Guides to the Identification of the Microinvertebrates of the Continental Waters of the World. 1996. ([100])

Key literature to identify Rotifer: Identification of rotifers should proceed with the use of taxonomic keys. Taxonomic identification, commonly

followed are as follows:

● E. McCauley, “The estimation of the abundance and biomass of mesozooplankton in samples,” in A

manual on methods for the assessment of secondary productivity in fresh waters, 1984, pp. 228–265.

([49])

● A. Ruttner-Kolisko, “Suggestions for biomass calculations of planktonic rotifers,” Arch. fur Hydrobiol.

Beihefte, vol. 21, pp. 71–76, 1977. ([76]) (species)

● K. W, Rotatoria. Die Radertiere Mitteleuropas Gebruder Borntraegerand. Berlin, 1978. ([101]) (species)

● W. T. Edmondson, “Rotifera,” Freshw. Biol., pp. 420–494, 1959. ([102])

● R. S. Stemberger, A guide to rotifers of the Laurentian Great Lakes, vol. 1. Environmental Monitoring and Support Laboratory, Office of Research and Development, US Environmental Protection Agency, 1979. ([103]). (genera)

● R. W. Pennak, “Fresh-water invertebrates of the United States 3rd Edition,” in Fresh-water invertebrates of the United States, 3rd ed., New York: Wiley New York, 1989. ([104])

● R. L. Wallace, “Rotifera,” eLS, 2001. ([105]) (freshwater Rotifera)

Other taxonomic identification literature ● H. Bottrell et al., “A review of some problems in mesozooplankton production studies,” Nor. J. Zool., vol.

24, pp. 419–456, 1976. ([48])

● Mekong River Commission and Environment Programme, “Identification Handbook of Freshwater Zooplankton of the Mekong River and its Tributaries Identification Handbook of Freshwater Zooplankton of the Mekong River and its Tributaries,” 2015. ([106])

● J. H. Thorp and A. P. Covich, Ecology and classification of North American freshwater invertebrates. Academic press, 2009. ([107])

● T. Nogrady, R. Pourriot, and H. Segers, “Rotifera 3. Notommatidae and Scaridiidae,” Guid. to Identif. Microinvertebrates Cont. Waters World 8.(H. Dumont, T. Nogrady, eds). SPB Acad. Publ. BV, 248 p., 1995. ([108]).

● H. Segers, “Rotifera 2. The Lecanidae (Monogononta),” Guid. to Identif. Microinvertebrates Cont. Waters World 6.(HJ Dumont, T. Nogrady, eds). SPB Acad. Publ. BV., 226 p., 1995. ([109])

● W. De Smet, “The Proalidae (Monogonanta),” in Guides to the identification of the macroinvertebrates of the continental waters of the world, H. Dumant, Ed. Amsterdam: SPB Academic publishing, 1995, pp. 1–102. ([110])

● W. De Smet and R. Pourriot, “The Dicranophoridae (Monogononta) and Ituridae (Monogononta),” in Rotifera Vol. 5, Guides to the identification of the macroinvertebrates of the continental waters of the

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world, H. Dumant, Ed. The Hague: SPB Academic publishing, 1997, p. 1–344. ([111]) ● K. Smith and C. H. Fernando, A guide to the freshwater calanoid and cyclopoid copepod Crustacea of

Ontario. Department of Biology, University of Waterloo, 1978. ([112]) ● M. D. Balcer, N. L. Korda, and S. I. Dodson, Zooplankton of the Great Lakes: a guide to the identification

and ecology of the common crustacean species. Univ of Wisconsin Press, 1984. ([113]) ● J. L. Brooks, “The systematics of North American Daphnia,” Mem. Connect. Acad. Art Sci., vol. 13, pp. 1–

180, 1957. ([114]) ● B. Dussart, Les Copépodes des eaux continentales d’Europe occidentale. 1. Calanoïdes et Harpacticoïdes.

Boubée, 1967. ([115]) ● B. Dussart, Les Copépodes des eaux continentales d’Europe occidentale...: Cyclopoïdes et biologie, vol. 2.

N. Boubée et Cie, 1969. ([116]) ● P. D. N. Hebert, “The Daphnia of North America: an illustrated fauna,” CD-ROM, Univ. Guelph, 1995.

([117]) ● N. N. Smirnov, Cladocera: the Chydorinae and Sayciinae (Chydoridae) of the world, vol. 11. SPB Academic

Pub., 1996. ([118]) ● J. W. Reid and H. Ueda, Copepoda: Cyclopoida: Genera Mesocyclops and Thermocyclops. Backhuys

Publishers, 2003. ([119]) ● A. Petrusek, F. Bastiansen, and K. Schwenk, “European Daphnia species (EDS)-taxonomic and genetic

keys.[Build 2006-01-12 beta],” CD-ROM Distrib. by authors. Dep. Ecol. Evol. JW Goethe-University, Frankfurt am Main, Ger. Dep. Ecol. Charles Univ. Prague, Czech Repub., 2005. ([120])

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Appendix E

1. E. McCauley, “The estimation of the abundance and biomass of mesozooplankton in samples,” in A

manual on methods for the assessment of secondary productivity in fresh waters, 1984, pp. 228–265 [49],

including the following meta-analysis:

a. H. J. Dumont, I. Van de Velde, and S. Dumont, “The dry weight estimate of biomass in a selection of Cladocera, Copepoda and Rotifera from the plankton, periphyton and benthos of continental waters,” Oecologia, vol. 19, no. 1, pp. 75–97, 1975. ([67])

b. H. Bottrell et al., “A review of some problems in mesozooplankton production studies,” Nor. J. Zool., vol. 24, pp. 419–456, 1976. ([48])

c. M. L. Pace and J. D. Orcutt, “The relative importance of protozoans, rotifers, and crustaceans in a freshwater mesozooplankton community,” Limnol. Oceanogr., vol. 26, no. 5, pp. 822–830, 1981. ([70])

d. R. A. Rosen, “Length-dry weight relationships of some freshwater mesozooplanktona,” J. Freshw. Ecol., vol. 1, no. 2, pp. 225–229, 1981. ([71])

e. G. Persson and G. Ekbohm, “Estimation of dry-weight in mesozooplankton populations - Methods applied to Crustacean populations from lakes in the Kuokkel area, Northern Sweden,” Arch. fur Hydrobiol., vol. 89, no. 1–2, pp. 225–246, 1980. ([68])

f. C. W. Burns, “Relation between filtering rate, temperature, and body size in four species of Daphnia,” Limnol. Oceanogr., vol. 14, no. 5, pp. 693–700, 1969. ([69])

g. M. J. Burgis, “Revised estimates for the biomass and production of mesozooplankton in Lake George, Uganda,” Freshw. Biol., vol. 4, no. 6, pp. 535–541, 1974. ([121])

h. T. R. Jacobsen and G. W. Comita, “Ammonia-nitrogen excretion in Daphnia pulex,” Hydrobiologia, vol. 51, no. 3, pp. 195–200, 1976. ([122])

i. E. Michaloudi, “Dry weights of the mesozooplankton of Lake Mikri Prespa (Macedonia, Greece),” Belgian J. Zool., vol. 135, no. 2, pp. 223–227, 2005. ([97])

2. L. A. Smock, “Relationships between body size and biomass of aquatic insects,” Freshw. Biol., vol. 10, no. 4, pp. 375–383, 1980. ([123])

3. D. A. Culver, M. M. Boucherle, D. J. Bean, and J. W. Fletcher, “Biomass of Freshwater Crustacean Zooplankton from Length–Weight Regressions,” Can. J. Fish. Aquat. Sci., vol. 42, no. 8, pp. 1380–1390, Aug. 1985. ([124])

4. J. Watkins, L. Rudstam, and K. Holeck, “Length-weight regressions for mesozooplankton biomass calculations – A review and a suggestion for standard equations.,” Cornell Biol. F. Station. Dep. Nat. Resour., 2011. ([125])

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14. Checklist for the next version

◻ Finalize Excel sheet on length-weight relationships.

◻ Add the Workflow (Section 9.1) if possible.

◻ Decide on a ‘specific conversion factor’ to estimate the mass as a carbon content, from

literature (if recommended).