Freshwater Recorders and Schemes Project

Sampling methodologies

This section is divided into the various habitats in which hypogean Crustacea can be found, with suggestions of potential sampling methods given for each.

THE SHALLOW INTERSTITIAL HABITAT

• The Bou-Rough Method
• The Karaman-Chappuis Method
• Other methods
  

SPRINGS

• Drift Nets
  

WELLS, PUMPS AND BOREHOLES

• Pit Wells
  • Trawl Net
  • Bait Traps
  • The FBA Well Sampler

• Other methods
  

CAVES AND MINES

• Subterranean rivers and streams
• Small pools, gour pools, dripping / trickling water on calcite slopes etc.
• Larger pools / underground lakes, sumps and flooded passages

PRESERVATION OF SAMPLES AND SPECIMENS

SAMPLE PROCESSING

Many of the nets mentioned in this section can be obtained from GB nets (now owned by Educational Field Equipment Ltd.) based in Bodmin (www.gbnets.info) or from various other suppliers of ecological field equipment. Less expensive small hand nets can also be obtained from aquarist shops. Choosing the size of the net mesh aperture requires a balance between capturing smaller sized specimens and maintaining a good flow of water through the net, preventing it from becoming clogged up by silt and debris. This is not usually a major problem in most subterranean environments where organic matter is low. The standard Environment Agency kick sampling / pond net with a mesh of 1mm is likely to retain most larger specimens, but small Niphargus glenniei, bathynellids and juveniles might be lost. A mesh of 250μm is likely to be more suitable for hypogean Crustacea sampling. 

Nets for use in sampling hypogean aquatic habitats. A: Drift net. B: Pipe Net (with draw-string). C: Aquarist’s hand net. D: Zooplankton trawl net (with lead weights). E: Hand net (with optional extended handle section, more than one can be fitted if required).

The best time to sample for hypogean crustaceans (and all groundwater fauna in general) is after heavy rainfall following a prolonged dry period. This tends to mobilise the groundwater fauna and wash them out of interstices and fissures in the rock strata into caves and wells etc. Repeat visits to wells and caves might sometimes be required, as in the past visits in different weather conditions have produced significantly different results.

THE SHALLOW INTERSTITIAL HABITAT

The shallow interstitial habitat consists of flooded alluvial (riverine) gravels and the hyporheic zone (zone of interaction between up-welling groundwater and down-welling river water) beneath watercourses. 

Being the most superficial of all the niphargids, Niphargus aquilex will occasionally turn up in samples of the epigean aquatic benthic macro-invertebrate fauna collected by kick sampling. N. aquilex can often be a significant component of the benthic fauna in previously dry winterbourne sections of rivers, after heavy rain has elevated the groundwater levels and re-started the surface flow. Over time N. aquilex will retreat back into the hyporheic zone as it faces predation and competition with Gammarus pulex and other epigean invertebrates. Recently Niphargus fontanus has also been identified in two Welsh Environment Agency kick samples, a species far less likely to be collected on the surface. Both N. aquilex and N. fontanus have been obtained by kick sampling an area of up-welling groundwater in a pond on chalk geology near Havant, Hampshire.

Bou-Rouch sampling equipment. A: Perforated zinced iron pipe. B: Piston pump attached to pipe. C: Protective striking cap. D: Hammer

Karaman (1935) and Chappuis (1942) both developed a method for sampling the fauna in the water beneath gravel banks at the margins of rivers and streams. It simply involves digging a hole into the gravel until the water level is reached and then either filtering the water as it flows into the hole, or collecting it with a jug and pouring it through a sieve or net.

Pospisil (1992) describes several other methods for sampling groundwater in shallow sediments. These include freeze coring, stand-pipe corers and the burying of artificial substrates. 

Freeze coring involves freezing the sediment around a stand-pipe and then removing the whole as a block, frozen fauna included.  

Stand-pipe corers remove small amounts of sediments from depths of about 1m but can yield uncertain results, due to their intrusive nature and the avoidance reactions of fauna.

Artificial substrates need to be left in-situ for several days or longer, enough time to be colonised by fauna from the surrounding habitat. Perhaps the best type is the Bishop Sampler, which consists of a box with moveable sides that is buried in the sediment with the sides open and filled with sediment. When it is due to be removed the sides are closed by wires and the sampler is dug out of the sediment. Simpler artificial substrates are of the Coleman and Hynes (1970) type, consisting of two perforated PVC tubes, one inside the other and filled with sediment.

The Noll spring sampler (diagram from Pospisil, P., 1992 ‘Sampling methods for groundwater animas of unconsolidated sediments’ in ‘The natural history of biospeleology’ Monografias Museo Nacional de Ciencas Naturales, Camach, A.I. (ed.))

Drift nets

The placement of drift nets at the point of issue can be effective in sampling springs. These simply consist of a net fixed in place and left to capture fauna as it is washed out of the ground. Camacho (1992) describes a method combining drift nets and the aid of cave divers for springs and resurgences that are accessible by humans. As the divers enter the spring the drift nets are anchored in place and removed when the divers exit. As the divers advance up the flooded gallery they agitate the silt on the bottom and walls of the gallery, dislodging fauna that are then washed out and into the nets. 

Pospisil (1992) describes a spring sampler developed by Noll (1939), consisting of a double funnel of bronze wire netting, fixed to a glass flask and sealed with a rubber ring. The device is dug into the mound of the spring’s issue point and removed after several hours.

These provide access to the deeper interstitial habitats and the phreatic groundwater. If a property is still known to be connected to a water supply from a well or borehole, one of the easiest sampling methods is to attach a pipe net (a simple net with a draw string or clamp) around the end of a tap or hydrant and leave it flowing for a while. Niphargus glenniei was first discovered in West Cornwall when specimens turned up in the water of a farm connected to a subterranean well.

Piezometers (of varying diameters from 2.5-20cm) are installed at many places for groundwater table and quality monitoring purposes. A tube can be inserted into a piezometer or borehole and connected to a pump; the flow generated by this is then passed through a sieve or net, or into a tank for holding sediments and fauna. A variety of pumps can be used, from hand-operated piston pumps to motorised pumps. However, the latter, whilst having a higher extraction power, can consequently damage delicate specimens.

Old accessible well shafts (pit wells) are fast disappearing as many properties are now on mains water and the old shafts, deemed to be unsafe, are either filled in or capped. However, such wells can provide good results and several methods for sampling these are described below.

Perhaps the easiest method is to lower a zooplankton net on a rope into the water at the bottom of the shaft several times and trawl it around the bottom, disturbing the sediment if possible. Nets are light (and consequently might require some weighting in order to sink properly) and easy to carry but there is the danger that they can become snagged on obstacles at the bottom of the well, such as large stones, branches and the remains of old piping etc. and also require space to be manoeuvred.

During the 1950s and 60s Glennie achieved very good results from wells using bait traps. Sometimes these consisted of nothing more complicated than jars lowered on a rope and left in-situ. A more effective trap can be made from plastic bottles by simply cutting off the top, inverting it and re-inserting it into the lower half and sealing the join with glue or heavy-duty tape.

Traps are probably best left in place for 24 hours and certainly not longer than 48 hours, as the bait itself can become a pollutant in the water if left for too long. There are various discussions on what constitutes good bait, although fish, liver and smelly (non-processed) cheeses seem to be the best. Some workers advocate fresh bait and some have achieved better results with slightly rotten bait.

A: Brass sleeve and gauze. B: Sampler bottles (in sampling position. C:Nylon air pipe with union and connector. D: Connector to attach bicycle or foot pump.

The former director of the FBA Hugh Gilson designed a simple device for effectively sampling wells. It consists of two plastic bottles, the upper inverted and connected to the lower, upright bottle by a length of copper tube, with a lead collar on the base of the lower bottle to maintain the sampler in an upright position when in use. A gauze mesh disc is fitted on the top of the upper bottle. An air-pipe, terminating in a ceramic aquarium diffuser enters the lower bottle at the shoulder. When in use, the sampler sits with the bottom of the lower bottle partially buried in the sediment at the bottom of the well. Air is pumped (by a suitable pump connected to a hose) into the lower bottle to agitate the sediment and wash out fauna, which is then washed into the upper bottle and trapped there by the mesh disc.

Pospisil (1992) describes two other methods for investigating deep interstitial habitats.  

The Double-packer Sampler is used in small diameter (2.5-7.5cm) monitoring wells (piezometers) which are perforated throughout their length. A sampling head is attached to a tube and fixed within the pipe at a set depth by inflatable tubes. Water is then pumped through the packerhead and tube by a piston or motorised pump. 

The second method involves inserting transparent pipes of 5cm diameter into the sediment. A small video camera, equipped with lights and a watertight casing and attached to a monitor is then introduced for in-situ observation of the sediment and its fauna. 

Caves especially contain a variety of aquatic habitats that can be sampled for hypogean fauna.

Many of the sampling methodologies used to sample the aquatic macro-invertebrate benthos in surface waters can also be used underground, although the main factor to consider is getting the equipment in and out of a difficult environment. These methods can include kick sampling, cylinder sampling, surber sampling, artificial substrates (both in the water and the sediment) and the use of drift nets. In allogenic streams (originating on the surface and flowing into cave systems) epigean invertebrates, washed in from the surface, will often be present, especially near the entrance. Autogenic streams (originating underground, within the cave system) are likely to produce better results with regards to hypogean fauna.

The Karaman-Chappuis and Bou-Rouch methods can be used to sample gravel beds at the margins of underground watercourses.

Small pools, gour pools, dripping / trickling water on calcite slopes etc.

These habitats are probably best sampled using manual searching, simply looking for fauna and collecting specimens using forceps or a pipette. It is recommended that entomological or feather-light forceps are used to avoid damaging specimens. Animals in thin / trickling films of water can be collected using small paint brushes, minimising damage. Care should be taken to avoid disturbing the sediment at the bottom of pools as this can quickly cloud the water and make manual searching impossible. This also means that manual searching is only effective in small pools in which the whole of the bottom can be viewed from the edges.   In slightly larger pools an aquarium hand-net (or similar), fitted with extended handle sections if required, can be used to sweep through the water and substrate at the bottom. This method can also be used to supplement the manual search in smaller pools if no taxa have been found, they might be hiding in the substrate at the bottom of the pool.

Care should be taken when sampling for any hypogean fauna not to over-collect and damage small, isolated populations. The populations seen in caves are really only the “tip of the iceberg”, compared to the populations in the surrounding rock strata and meso-cavernous spaces, but as with all natural history study, only collect the minimum number of specimens required for identification or none at all.
Most hypogean Crustacea seem to prefer pools with fine silt at the bottom, although they are occasionally seen in crystal-lined gour pools. Whilst feeding, Niphargus species often leave small trails on the surface of silt and these can be good indicators that taxa are present. However, due to the humidity in caves, it is not unusual for them to exit the water and migrate between pools or back into fissures in the surrounding rock over damp sediments and rocks.

A lot of water can enter cave systems through cracks and fissures in the roof and walls. After heavy rainfall these can become quite active and might contain fauna washed out of the surrounding rock strata. Jars, or small nets fixed in place beneath the fissures, can serve to collect specimens. Carmacho (1992) states that such diffuse filtrations can be sampled by placing a mesh, slightly detached from the ceiling, and set in such a way that it channels the water towards a group of funnels with a net to retain fauna. Such an apparatus could be placed over a group of fissures and left as a semi-permanent fixture, as only the final net would need to be checked and regularly emptied. 

Larger pools / underground lakes, sumps and flooded passages

Manual searching or the use of nets, especially those fitted with additional handle sections for extra length, can be used to sample the margins of such waterbodies but will obviously not sample the deeper sections. 

The deeper sections can be sampled using a trawl net (zooplankton net) attached to a length of rope. The net will probably need to be weighted in order to be thrown from the edge of the waterbody effectively and sink to the bottom. There is also the risk that the net will become snagged on submerged rocks.
Bait traps can also be used in the larger water bodies, as discussed in the section on pit-wells. However, the use of bait traps is not recommended in caves as they involve two visits to place and retrieve and the bait can cause organic contamination of the water. Also there is the risk that a sudden flood can displace traps so that they cannot be recovered. In the cave environment, where food is scarce, the bait will then become a long-lasting focus of attraction which could ruin small, localised populations of hypogean fauna. Predators might also enter the traps and devour the fauna that has gathered.
Artificial substrates are another possible method but again would involve two visits to place and retrieve and could amount to a lot of encumbrance to carry, along with any other caving equipment that might be needed.

Camacho (1992) mentions the use of manual searching in sumps by cave divers, collecting fauna using modified nets. She also mentions a manual aspirator which works with compressed air and is very effective at collecting fauna from submerged cracks and clay walls, although it can cause considerable damage to the delicate specimens.

PRESERVATION OF SAMPLES AND SPECIMENS

The two main chemicals used for the preservation of aquatic fauna are formalin solution and alcohol solution. The former, usually used as a 5% solution, is good at fixing specimens, especially delicate structures such as gills and legs. However, it is also very acidic and needs to be buffered, as over time it will dissolve calcareous structures, such as snail shells and crustacean exo-skeletons. Formalin is also highly toxic and carcinogenic.

Alcohol is the safer option and is normally used as a 70% solution, although this can damage specimens and make them brittle. An initial killing solution of 40% is best, with the specimens later placed in a 70% solution for long-term storage if required. The addition of a small amount of glycerol will also prevent complete dehydration of the specimens should the tubes become damaged. If specimens are required for DNA analysis then they must be preserved in 100% alcohol (preferably ethanol, although bear in mind that ethanol cannot be made 100% as it always contains 6% water).

The most commonly used alcohol solution is industrial methylated spirits (IMS), although this will require a licence from HM Customs and Excise to obtain. Other alcohols used include ethanol and isopropanol. If the solution is required for short-term storage, (i.e. for sending on to an expert for identification / confirmation) then standard, purple-dyed methylated spirits (as used in camping stoves etc.), or even a strong spirit such as vodka or rum can be used. However, note that the purple methylated spirits should be used neat and preferably only as a last resort, since it forms a white flocculent precipitate when added to water.

Most samples from underground environments should contain little organic matter, so separation of the fauna from a predominately mineral substrate should not be too difficult. They are also not likely to contain large snails and cased caddis larvae with stony cases, as found in epigean samples, which will probably be too heavy to separate out easily. 

The easiest method is by decantation and sieving. The sample (or fractions of larger samples) is washed in a large tray and the sediment agitated so that fauna float to the surface. The supernatant is then poured through a sieve and fractions of the sieved matter are then sorted through on another tray with sufficient illumination and preferably some sort of low level magnification. Specimens are then picked out for further examination under a binocular microscope. It is wise to check the substrate left in the first tray, to make sure that no fauna has been missed.

Another method is to use the modified flotation technique developed by Anderson (1959), involving the introduction of sugar solution (Approx. 2.5lbs granulated sugar per gallon of solution) to the sample, which will cause organic matter to float to the surface, due to differences in the specific gravities of the organic matter and the mineral substrate.

A third method is to use staining, with agents such as 1% eosine, which stains organic matter pink and hence makes it easier to identify. This staining requires at least 24 hours and the sample will need to be refrigerated to prevent deterioration.

Sometimes it is easier to sort samples live, as the movement of the animals in a tray can make them easier to spot and pick out. It is preferable to do this in a dark-coloured tray, against which the white hypogean organisms will stand out better. However, small specimens such as juvenile Niphargus and especially bathynellids are likely to be missed.