Detailed protocols for sampling

%moisture and % ash free dry mass (AFDM)

In the laboratory, sieve samples (2mm). Samples are then weighed, dried at 60°C for 24 h, and re-weighed, to calculate water content (%), and combusted at 550°C for 4 h to estimate AFDM.

In total, these samples comprise:

Flowing phase: 1 sediment sample (6 pooled subsamples) from channel and 1 soil sample (6 pooled subsamples) from riparian habitat

Non-flowing phase: 1 sediment sample (6 pooled subsamples) from channel and 1 soil sample (6 pooled subsamples) from riparian habitat

Total number of samples: 8 = 2 reaches (1 perennial + 1 intermittent) * 2 hydrological phases (flowing + non-flowing) * 2 habitats

Determine % water content and %AFDM for each for the 8 samples.

Surber samples

During the flowing phase, collect 2 Surber samples (sample area 0.1 m2, mesh size: 250 µm) from 2 riffles and 1 Surber samples from each of 2 pools in the intermittent reach, and the same (another 4 samples) in the perennial reach. During the non-flowing phase, if any disconnected pools are present, sample 2 pools if you can choose, or from 1 if only 1 is left. If the pools are too small, i.e., < 1×1 m, disregard them. Surber samples are taken by vigorous manual disturbance of the substrate within the Surber quadrat for one minute. To generate flow, some agitation by hand is needed to transfer disturbed invertebrates into the net and have an effective sampling in lentic conditions. Use a comparable method in the riffle and pool habitats in the perennial reach For each sample, empty the contents of the net into a container and preserve in a 70% ethanol. Nets should be thoroughly checked and any invertebrates remaining on the net should be removed and put in the sample container.

Invertebrates will later be identified and counted by each participant team (in your own lab) to the lowest possible identification level. Invertebrate data, including taxon list and counts, along with information on the hydrological phase, should then be computerized and sent to the core team of the 1000IRP project.

Pitfall traps

For each reach (perennial, intermittent) and each phase (flowing, non-flowing), establish 6 spots, located a certain distance apart from each other (i.e. 2 ×mean channel width) and perpendicular to the thalweg channel. At each spot, sample two habitat types (channel and riparian). The channel habitat is defined as (i) the center of the dry stream channel in the intermittent reach during the dry phase, and as (ii) the shoreline of the stream channel (i.e. the exposed stream sediments located a few 10-20 cm centimeters from the water’s edge) in the intermittent reach during the dry phase and in the perennial reach during both phases. The riparian habitat in both types of reaches and for both hydrological phases is defined as the area located from the edge of the high-water channel and bankfull to the edge of upland, characterized by the presence of a distinctive riparian vegetation and substrate type. Within each habitat type at each spot, deploy 1 pitfall trap. Distribute shoreline and riparian spots evenly on both sides of the channel (i.e. 3 on right side and 3 on left side). Pitfall traps are plastic containers (height: approx. 80 mm, diameter: approx. 80mm) inserted into the sediments and filled to 3/4 with glycol (cooling liquid for cars, very toxic). Record the exact height and diameter of the containers used. Position a plastic cover over each trap to prevent rain, falling leaf litter and other debris from blocking the trap. Set the traps for one week (min. 5 days, max 15 days). Pool the contents of all collected pitfall traps from each habitat at one site, transfer the collected specimens into plastic containers filled with 70% ethanol, and store until further analysis. Record the number of pitfall traps pooled for each habitat, in case some were lost during sampling (e.g. destroyed by animals).

In total, these samples comprise:

Flowing phase: 1 sample (6 pooled pitfall traps) from channel and 1 sample (6 pooled pitfall traps) from riparian habitat

Non-flowing phase: 1 sample (6 pooled pitfall traps) from channel and 1 sample (6 pooled pitfall traps) from riparian habitat

Total number of samples: 8 = 2 reaches (1 perennial + 1 intermittent) * 2 hydrological phases (flowing + non-flowing) * 2 habitats

In the lab, identify the sampled terrestrial invertebrates to the lowest taxonomic level possible. See Corti et al. 2013 for details. Invertebrate data, including a taxa list and counts, along with information on the hydrological phase, should then be computerized and sent to the core team of the 1000IRP project.

eDNA sampling from the water column

During the flowing and non-flowing phases, collect eDNA in 2 pools (for each reach) by filtering water through a 0.22 µm filter using Sterivex cartridges (http://www.merckmillipore.com/FR/fr/product/Sterivex-GP-Pressure-Filter-Unit,MM_NF-SVGPL10RC). Filtration should be done in the field using a peristaltic pump or a 50-mL sterile syringe until the filter clogs, and the duration of the filtration procedure (in seconds) and volume filtered (in mL) recorded. At the end of the filtration step, remove excess water from the Sterivex cartridge by pumping air through the filter. The volume of water filtered and the duration of the filtration procedure are mandatory information. Label (using the 1000IRP recommendations explained in section 3.6) your Sterivex cartridge and place the cartridge in a sealed zip-lock bag with the same label.  Store filtered water samples at -20°C until shipping (see below).

In total, the required eDNA samples comprise:

Flowing phase: 2 samples from pools

Non-flowing phase: 2 samples from pools

Total number of samples: 8 = 2 reaches (1 perennial + 1 intermittent) * 2 hydrological phases (flowing + non-flowing) * 2 pools

Soil and sediment samples for eDNA analyses

Collect soil and sediment samples (2 L in volume) in 3 locations within a circular sampling area with 1 m radius and at depths of 0-10 cm using a shovel. For the flowing and non-flowing phases, 4 sediment samples are collected from the riverbed (2 from pools and 2 from riffles) and 4 soil samples are collected in the adjacent riparian zone (two on each side of the river channel and < 20 m from the riverbed) at each site for eDNA analyses. Avoid cross contamination of samples during the sieving procedure. Wear gloves and clean the material used for soil/sediment sampling with ethanol (shovel, sieves) between each sample. In the laboratory, sieve soil or sediment samples at 2 mm. Take 2 subsamples of about 15 g from the sieved and homogenized 2 L sample and store them at -20°C in 2 sterile 50 mL labelled Falcon tubes. For each sample, send 1 Falcon tube to Irstea and retain the other (replicate subsample) at -20°C as a backup.

In total, the required eDNA samples comprise:

Flowing phase:

  • Riverbed: 2 sediment samples from pools, 2 sediment samples from riffles
  • Riparian zone: 4 soil samples (two on each side of the river channel and < 20 m from the riverbed)

Non-flowing phase:

  • Riverbed: 2 dry sediment samples from pools, 2 dry sediment samples from riffles
  • Riparian zone: 4 soil samples (two on each side of the river channel and < 20m from the riverbed)

Total number of samples: 32 = 2 reaches (1 perennial + 1 intermittent) * 2 hydrological phases (flowing + non-flowing) * 8 (2 pools, 2 riffles, 4 riparian)

Invertebrate seedbank:

The riparian habitat is defined as the area located from the edge of the high-water channel and bankfull to the edge of upland, characterized by the presence of a distinctive riparian vegetation and substrate type. During the non-flowing phase, each sediment sample (2.5 L in volume, sieved to retain the fraction < 5 mm) is collected at depths of 0-10 cm using a shovel. Estimate the time since the last flowing period (even if with some uncertainty, i.e. ± 1 week). Samples are placed in individual plastic buckets of 10 L, returned to the laboratory, and if possible, placed in an environmental chamber at room temperature (20°C) with a day-night cycle corresponding to the conditions at the sampling reach. Then, 5 L of tap water, previously dechlorinated by standing it for 24 h in the lab, is added to each bucket. After water is added, each bucket is gently shaken, and the surface water poured through a 250-µm metal sieve. This initial sample is used to collect any living aquatic or terrestrial invertebrates and predators that may prey on viable eggs and is preserved with 70% isopropyl alcohol. Each bucket is then refilled with tap water previously dechlorinated to an equivalent 5-L level, an air-stone is placed on the sediment surface to keep the water oxygenated, and each bucket is covered with a plastic screen (1-mm mesh) to keep any emerging insects from escaping. The total incubation/inundation period is 21 d, as a compromise between time for invertebrates to emerge and preventing anoxia to develop. After 21 d, sediments in each bucket are gently elutriated 5 times with each elutriate decanted through a 250 µm mesh sieve. The material retained on the sieve is preserved with 70% isopropyl alcohol for later analysis. All invertebrates are handpicked and identified to the lowest practical level and counted. See more details in Larned et al. 2007, Datry et al. 2012. Send the list of invertebrates and eggs counted and identified to the core team of the 1000IRP project, along with information on the hydrological phases.

Photo Traps:

During the whole study period, each sampling reach is monitored using a photo-trap. This device records animal movements when they cross the ‘field’ of the camera, allowing their identification from the recordings later on. To do so, a device should be placed in a nearby tree, so it can cover a maximum of the riverbed, including when dry. The frequency of downloading the images recorded is device-dependent, so this is up to each participant. For more information, see Findlay et al. 2017. After the study period send to the core team of the 1000IRP project the list of animals recorded along with information on number of individuals and in which phase and time the animals appeared.

Tracks:

For each phase, identify 2 habitat types: dry riverbed and riparian zone habitats, separated from each other by approximately 100 m. In each habitat, select 3 stations each comprising 1 circle of 0.7 m in diameter, which will be covered with a 2-5 mm layer of smoothed white marble or fine sand dust. Try to place one station in the range of the phototrap installed (see 5.7). In case of rain, the layer may need to be reset. Stations are checked every 2 days to observe the tracks of terrestrial vertebrates (optimum would be 5 visits per station). After documenting the number and the direction of tracks, the dust surface is resmoothed.

To document tracks:

-take photos

-note the direction of the tracks: upstream, downstream, perpendicular to the river on the right, perpendicular to the river on the left, no clear direction

-classify the animal according to its taxonomic group: micromammals (Orders Rodentia and Insectivora), lagomorphs (Order Lagomorpha), marsupials (Diprotodontia),  carnivores (Order Carnivora), ungulates (Order Artiodactyla), reptiles (Class Reptilia), and birds (Class Aves)

– use the latin name for the species/genus, whenever possible.

For more information, see Sánchez-Montoya et al. 2016. After the study period, send the list of animal tracks identified to the core team of the 1000IRP project, along with information on number of individuals and in which phase tracks were recorded.