FunAqua sampling protocol: standardized methods for fungal diversity assessment in aquatic ecosystems

FunAqua sampling protocols Version: 1.0 | DOI: 10.5281/zenodo.17750254 1 INTRODUCTION Fungi in aquatic ecosystems play diverse and ecologically significant roles; however, they remain understudied, and their global diversity is poorly understood compared to other aquatic microorganisms and terrestrial fungi. While fungal diversity has been reported in various aquatic environments, inconsistencies in sampling strategies and molecular identification methods have hindered a comprehensive understanding of dominant taxa, overall diversity, and the ecological processes shaping fungal communities. The FunAqua project aims to address these gaps by systematically investigating fungal diversity in aquatic ecosystems and its environmental drivers. Our objective is to characterize fungal birodiversity across local and global gradients and explore their connections with soil and leaf-associated communities. This protocol provides standardized procedures for collecting water and sediment samples from lakes, rivers, and marine systems. Samples will be processed through DNA extraction and high-throughput sequencing of ribosomal RNA gene regions (SSU and ITS), ensuring comparability across international collaborators. By harmonizing sampling methods, FunAqua aims to advance knowledge of aquatic fungal ecology and deliver insights into their roles in ecosystem functioning. 2 BASIC SAMPLING PROTOCOL 2.1 OVERALL REQUIREMENTS Required fieldwork material: 1. Global Positioning System (GPS) receiver 2. Sterile 0.5-1 L bottle(s) (preferentially polyethylene, PE or glass) for water samples. 3. Two sterile 50 mL tubes (e.g. centrifuge tubes, polypropylene) or plastic bottles for sediment samples. 4. Waterproof marker pen. 5. Powder-free disposable gloves 6. Recommended: a coolbox with ice or ice blocks to keep the samples in dark and cold. 7. If availble, Sterivex filters - cat number SVGPL10RC (protocol DOI: 10.5281/zenodo.7875535.) Required laboratory material: 1. Two membrane filters, pore size 0.2 μm, diameter 47 mm. Preferential types of filter membrane: mixed cellulose ester (MCE; ME), cellulose nitrate (CN), polyestersulphane (PES), or Polycarbonate Track Etched (PCTE) 2. Filtration device for 47 mm diameter filters 3. Vacuum pump 4. Tubes - preferably 2 ml screw cap tubes with o-ring. If these are not available, othersized screw cap tubes are an option for storing and transporting filters. 2-ml snap-lock tubes can be used, but are not recommended 5. 50-100 ml tube/bottle for water samples chemical analysis 6. 90-96% ethanol 7. Powder-free disposable gloves 8. Forceps 9. Transparent tape to cover all labels 10. Parafilm or insulating tape to cover the screw cap (to avoid leaking during transport) Supporting information to be recorded in the field/lab protocol: 1. Name and contacts of the collector 2. Name of the waterbody 3. GPS coordinates in WGS84 system: preferred format DD.DDDDD North, DD.DDDDD East 4. Sampling date: preferred format YYYY-MM-DD 5. Sampling depth (m) 6. Filtration volume (ml) for water samples 7. Filter type for water samples 2.2 IN THE FIELD SAMPLING - WATER 1. Label the bottle(s). 2. Walk carefully (try to prevent bottom sediments from rising) to a water depth of at least 40-50 cm. 3. Record the GPS coordinates of the sampling site, mark them in the notebook. 4. Put on disposable gloves (non-powdered). 5. Rinse the 1 L bottle(s) 3 times with surface water. 6. Place the bottle(s) approximately 20-30 cm below the water surface and fill with water. 7. Keep the samples in the dark. If possible, place the filled bottle in the cooler box for transportation. 8. NB! Sterivex filtering is recommended to be performed immediately after sampling. SAMPLING – SEDIMENT 1. Label two 50-ml tubes. 2. Walk to the water depth of 40-50 cm – preferably the same point as for water samples. * If not, record the GPS coordinates of the sampling site and mark these in the notebook. 3. Put on disposable gloves (non-powdered). 4. Rinse the two labeled 50 ml tubes 3 times with surface water. 5. Fill both tubes to half of their volume with sediment by pushing the tubes into the sediment layer, preferably at a sediment depth of 5-10 cm. 6. Pour off water and surplus material and fill the other half of the tube with 90-96% ethanol ASAP. Mix. 7. Keep the samples in a dark and cool place and transport them to the lab as soon as possible. 2.3 IN THE LABORATORY WATER SAMPLE FILTRATION AND FIXATION 1. Mix the sampled water before filtration by turning each bottle carefully several times upside down. 2. Optional: Collect 50-100 mL of water in a separate bottle (for chemical analysis) and store at -20 °C. 3. Filter a certain amount of water in 2 replicates through a 0.2 μm membrane filter, diameter 47 mm (preferential types of filter membrane are shown in the section required laboratory materials). Expected filtration volumes: 50–100 ml for hypertrophic water, 100-250 ml for eutrophic water and up to 1000 ml for oligotrophic water. 4. Write the filtration volume on the tube label and in your notebook. 5. For each filter, fill screw cap tubes with 90-96 % ethanol (use the amount of EtOH to ensure that the filter is submerged in it). Use the forceps to fold the filters on the filtering device as you see in Figure 1 below, and place them in separate tubes filled with ethanol. 6. Secure the cap with parafilm or insulation tape to prevent leaking. 7. Label all tubes 8. Cover the labels with transparent tape to protect them from ethanol leakage. 9. Store samples at -20 °C. 10. Ship sample #1 and the 50-100 ml of (optionally collected) water. As a backup, store sample #2 Figure 1. Filter folding SEDIMENT SAMPLE FIXATION 1. If samples were not fixed immediately after sampling, add 25-30 ml of 90-96% ethanol (fill another half of the 50 ml tube) to both of the half-filled sediment sample tubes. 2. Secure the cap with parafilm or insulation tape to prevent leaking. 3. Label all tubes as shown above, including the date, name of the waterbody, and the sample ID. 4. Cover the labels with transparent tape to protect them from ethanol leakage. 5. Store samples at -20 °C. 6. Ship sample #1. As a backup, store sample #2 3 ADVANCED SAMPLING PROTOCOL 3.1 OVERALL REQUIREMENTS Required fieldwork material: 1. Global Positioning System (GPS) receiver 2. Boat 3. Instrument(s) for measuring water temperature, DO, depth and other parameters available (pH, conductivity, etc) 4. Water sampler 5. Secchi disc 6. Optional: sediment corer 7. 3-6 sterile 1 L bottles (preferably polyethylene, PE, or glass) for water samples. 8. Two sterile 50 mL tubes (e.g., centrifuge tubes) or bottles for sediment samples. 9. Powder-free disposable gloves 10. Recommended: a cooler box with ice or ice blocks to keep the samples in a dark and cold place. If avialble, Sterivex filters - cat number SVGPL10RC (protocol DOI: 10.5281/zenodo.7875535.) Laboratory material: 1. 6 membrane filters, pore size 0.2 μm, diameter 47 mm. Preferential types of filter membrane: mixed cellulose ester (MCE; ME), cellulose nitrate (CN), polyestersulphane (PES), or Polycarbonate Track Etched (PCTE). * If available, use closed Sterivex filters 2. Filtration device for 47 mm diameter filters 3. Vacuum pump 4. Tubes-preferably 2 ml screw cap tubes with o-ring. If these are not available, other-sized screw cap tubes are an option for storing and transporting filters. 5. 90-96% ethanol 6. 50-100 ml tube/bottle for water samples chemical analysis 7. Powder-free disposable gloves 8. Forceps 9. Transparent tape to cover all labels. 10. Parafilm or insulating tape to cover the screw cap (to avoid leaking during the transport) Supporting information to be recorded in the field/lab protocol: 1. Name and contacts of the collector 2. Name of the waterbody 3. GPS coordinates in WGS84 system: preferred format DD.DDDDD North, DD.DDDDD East 4. Sampling date: preferred format YYYY-MM-DD 5. Overall water depth 6. If your waterbody is stratified, depth range of the thermocline (defined as the depth interval at which the rate of decrease of temperature with increase of depth is the largest) 7. Sampling depth (m) 8. Filtration volume (ml) for water samples 9. Filter type for water samples 10. Water temperature, if possible in every 0.5 m 11. Water pH and other abiotic parameters available 12. Water dissolved oxygen 13. Secchi depth 14. Water Ctotal, Ntotal, Ptotal, K, Mg, Ca concentrations OR send 50-100 ml of water to us for analysis 3.2 IN THE FIELD SAMPLING - WATER 1. Use a boat for sampling. 2. Identify the correct sampling point in the water body. Preferentially use the historical sampling point on which long-term records are based, or as close to the central point (if there is no history of sampling) as possible. 3. Label the bottles 4. Record the GPS coordinates of the sampling site, and mark these in the notebook. 5. Measure the abiotic parameters like temperature, DO, secchi depth, and other parameters available (pH, conductivity, etc) at 0.5-1 m intervals, record and note the values. 6. Define the depth range of the thermocline (the depth interval at which the rate of decrease of temperature with increase of depth is the largest) 7. Put on disposable gloves (non-powdered). 8. Rinse the labeled bottles 3 times with surface water. 9. From stratified waterbodies, take water samples from 3 layers: 1. 30 cm below the water surface, 2. from the thermocline 3. 1 m from the bottom of the water body (or as deep as possible in case of deep water bodies) From non-stratified (shallow) waters, take water samples from 2 layers: 1. 30 cm below the water surface, 2. 1 m from the bottom of the water body (or as deep as possible in case of deep water bodies) 10. Keep the samples in a cool and dark place and transport them to the lab as soon as possible. 11. NB! Sterivex filtering is recommended to be performed immediately after sampling (Protocol DOI: 10.5281/zenodo.7875535.) SAMPLING – SEDIMENT A. Sampling with a sediment corer 1. If possible, use the same sampling point as for water samples; if not, record the GPS coordinates of the sampling site and mark these in the notebook. 2. Put on disposable gloves (non-powdered). 3. Rinse 2 labeled 50 ml tubes or bottles 3 times with surface water. 4. Use a sediment corer to take the top 15 cm of sediment. 5. Fill two 50 mL tubes to half of their volume with the upper 0-10 cm of the sediment layer. 6. Pour off water and surplus material. 7. Fill the other half of the tube with 90-96% ethanol as soon as possible. 8. Keep the samples in a dark and cool if possible B. Sampling without a sediment corer - follow the basic protocol 3.3 IN THE LABORATORY WATER SAMPLE FILTRATION AND FIXATION 1. Mix the sampled water before filtration by turning each bottle carefully several times upside down. 2. Collect 50-100 ml of water from each sampled layer in a separate tube/bottle (for chemical analysis) and store at -20 °C. 3. Filter a certain amount of water in two replicates per sampled water layer through a 0.2 μm membrane filter, diameter 47 mm (preferential types of filter membrane are shown in the section required laboratory materials). Expected filtration volumes: 50-100 ml for hypertrophic water, 100-250 ml for eutrophic water, and up to 1000 ml for oligotrophic water. 4. Write the filtration volume on the tube label and in your notebook. 5. For each filter, fill screw cap tubes with 90-96 % ethanol (For 2 ml tubes, use 1.5 ml EtOH, for other volume tubes, use the amount of EtOH in which the filter is submerged). Use the forceps to fold the filters on the filtering device as you see in Figure 1 above, and place them in separate tubes filled with ethanol. 6. Secure the cap with parafilm or insulation tape to prevent leaking. 7. Label all tubes, including the name of the waterbody, the sample number, and the sampling depth. 8. Cover the labels with transparent tape to protect them from ethanol leakage. 9. Store samples at -20 °C. 10. Ship sample #1 (and 50-100 ml water if chemical analysis is collected). As a backup, store sample #2 SEDIMENT SAMPLES 1. Secure the cap with parafilm or insulation tape to prevent leaking. 2. Label all tubes, including the name of the waterbody, date, the sample number 3. Cover the labels with transparent tape to protect them from ethanol leakage. 4. Store samples at -20 °C. 5. Ship sample #1. As a backup, store sample #2 NOTE for GPS. Use a GPS device to record the coordinates of your sampling point. If water and sediment sampling points differ, record their coordinates separately. The commonly used datum in Europe is WGS84; therefore, all GPS units should be configured to this standard before fieldwork. Confirm that the WGS84 datum is active when the GPS is turned on prior to data collection. If the GPS cannot be set to WGS84, note the datum used on the data forms for later conversion. Important: Check the GPS batteries before departure. FunAqua Project | Version 1.0 | DOI: [placeholder] 4 CONTACT For questions or collaboration inquiries, please contact: Kristel Panksep, PhD FunAqua Project Coordinator Researcher in Molecular Ecology Estonian University of Life Sciences, Chair of Hydrobiology and Fisheries University of Tartu, Institute of Technology Swedish University of Agricultural Sciences, Department of Aquatic Resources (SLU Aqua) Email: [email protected]