2.0 Materials and Methods 2.1 Water Sampling Beacause of the close association between riparian and in-stream ecosystems (Blühdorn, 1997) and the likelihood that aquatic macroinvertebrates form at least part of the diet of E. yabbimunna (Suter & Richardson, 1977), a decision was made to evaluate the water quality in Shorewell Creek as part of this project. 2.1 The Study Sites Four sites were chosen along the length of the creek so as to reflect changes in water quality from the upper to lower catchment (see Fig. 1). Water was monitored on a weekly basis. Site W1 was located at the shallow end of the water storage in the headwaters of Shorewell Creek (see Fig. 2). An active tip site is located in an adjacent catchment. Although it was recognised that a dam is not an ideal site from which to sample streamwater it was decided to proceed because of the need to obtain a sample prior to any possible inputs into the stream. The vegetation at the site consisted of introduced grasses as well as plants which have escaped from domestic gardens. The water level fluctuated quite rapidly, depending on rainfall. Click to Enlarge Fig. 2: Water Sampling Site No. W1 at the Headwaters of Shorewell Creek. Site W2 was located in an area where the stream flowed through a relatively steep-sided valley which sloped down from housing estates on either side. The stream width was quite narrow and there were several large stormwater outlets discharging into the creek. The vegetation consisted mainly of willows (Salix spp.), eucalypts, blackwoods and man ferns (see Fig. 3). Click to Enlarge Fig. 3 : Water Sampling Site No. W2 in an Urban Environment on Shorewell Creek. Site W3 was located on the downstream side of a disused tip (see Fig. 4). Water is transported in buried pipes for a distance of approximately ¾ km through the old tip before emerging at site W3 iron-stained and with a very strong metallic odour. The vegetation here consisted of long grasses, blackwoods and man ferns. Click to Enlarge Fig. 4: Water Sampling Site W3 Downstream from the Disused Tip Site W4 was located in the Burnie Park (see Fig. 5). Here the stream flows through a man made rock channel with steep valley walls on either side. There were stormwater inputs from streets located on the ridgetops. A waterfall and duckpond are located upstream. Vegetation consised mainly of native species including blackwoods, eucalypts and manferns. Click to Enlarge Fig. 5: Water Sampling Site W4 at Burnie Park in the Lower Catchment Five physicochemical parameters were assessed, while coliforms and aquatic macroinvertebrates were used as biological indicators of water qualilty. Water sampling methods were closely based on those outlined in Waterwatch Tasmania (1996). 2.1.1 pH pH is a measure of acidity or alkalinity on a scale of 0-14. Between pH 7 and pH 0 a solution becomes increasingly acid, while from pH 7 to pH 14 a solution becomes more alklaline. The pH of stream water usually lies between pH 6.5 and pH 8.2 (Waterwatch Tasmania, 1996). Any major fluctuations in stream pH levels will result in the loss of more sensitive species of plant and animals. ANZECC (1992) recommends that pH levels be maintained at between 6.5 and 9.0 to ensure the protection of aquatic ecosystems. For this project pH was measured using a pHydrion Lo Ion Test Kit. pH paper which was sensitive to near neutral pH levels was shaken with a water sample and the pH was determined by comparing the resulting colour change with the colour chart provided. Waterwatch Tasmania considers this method to be an appropriate substitute for pH meters for the field monitoring of stream pH (W. Butler, Co-ordinator, Five Rivers Waterwatch Group, pers. comm.). 2.1.2 Alkalinity Alkalinity is a measure of water's resistance to changes in pH (GLOBE Program, 1997). Alkalinity in streamwater may be generated as water runs through rocks containing calcium carbonate (CaCO3) (e.g. limestone, dolomite) or where runoff contains appreciable quantities of CaCO3 such as where lime has been used for agricultural purposes. Alkalinity can be expressed in several forms, but one of the most common is as mg/L of CaCO3. GLOBE Program (1997) suggests that alkalinity levels of >100 mg/L CaCO3 are needed to absorb pH changes in streamwater. In areas where little calcium carbonate is present in the environment streamwater may contain low levels of alkalinity. For this project alkalinity was measured using a La Motte Alkalinity Field Test Kit. A water sample was titrated against sulphuric acid (H2SO4) using bromocresol green-methyl red indicator. Alkalinity was read as mg/L CaCO3 directly from the Direct Reading Titrator supplied with the kit. 2.1.3 Temperature Many physical, chemical and biological characteristics of a stream are directly affected by temperature. Waterwatch Tasmania (1996) lists several factors which influence the temperature of water in streams. These include: depth of the water the season time of day amount of streamside vegetation warm urban runoff from footpaths processes which increase turbidity such as soil erosion, road construction, cattle in streams discharge from reservoirs/dams which lower natural temperature levels Water temperature will alter the amount of oxygen dissolved in the water, affect the rate of photosynthesis of aquatic plants and influence the metabolic rate of fish and other aquatic animals (Waterewatch, Tasmania). Organisms can tolerate slow changes in temperature. ANZECC (1992) recommends <2% increase in temperature over 24 hours. Temperature was measured using a La Motte 0-50"C thermometer. The thermometer was placed into a water sample immediately after the sample was removed from the stream and the temperature recorded to the nearest 0.1"C after a period of 1 minute. 2.1.4 Turbidity Turbidity is the cloudiness of water and is the result of suspended material in the water (Waterwatch Tasmania, 1996). Stream turbidity most often results from soil erosion in the catchment or algal growth in the water (Jolly, et. al., 1996). This suspended material decreases the passage of light through the water which can limit plant growth and in turn affect the fish and invertebrate communities which feed on and live in the plants. Turbidity is commonly measured in Nephelometric Turbidity Units (NTU) and Jolly, et. al. (1996) report that 0-25 NTU is optimum for most aquatic life. Turbidity was measured with a "Waterwatch" Turbidity Tube. A sample of water was poured into the tube until the black pattern at the bottom of the tube was just visible. Turbidity was then read from the scale on the side of the tube. 2.1.5 Dissolved Oxygen Dissolved oxygen (DO) refers to oxygen molecules that are dissolved in water (Allaby, 1983). The presence of adequate levels of dissolved oxygen are essential for the respiration of aquatic organisms. The amount of oxygen that can dissolve in water decreases with increasing water temperature, so that at 20°C water is 100% saturated when oxygen levels reach 6.5 mg/L compared to 10 mg/L at 0°C. DO levels are also higher in turbulent water because of the constant exposure of the water to air. Bacterial decay will deplete DO levels. For this project, DO was measured using a La Motte Dissolved Oxygen Field test Kit. The initial concentration of DO in the sample was "fixed" using solutions of manganous sulphate, alkaline potassium iodide azide and sulphuric acid. The "fixed" solution was then titrated against 0.025 N sodium thiosulphate using starch as an indicator. The concentration of DO in mg/L was read directly from the Direct Reading Titrator supplied with the kit. 2.1.6 Coliform Bacteria The coliform group of bacteria grow naturally in the intestines of warm blooded animals. Coliforms are therefore present in large numbers in excrement (faecal matter). Their presence in waterways indicates that excrement from birds, animals or humans has recently polluted the water and that pathogens (organisms capable of causing disease) may also be present. Total coliforms were assessed using the Coliscan Easygel method. Water samples were collected in sterile containers and transported back to the laboratory. A sterile pipette was used to transfer a 2 mL sample into the supplied bottle of nutrient. After gentle swirling the nutrient-sample mix was tranferred to a petri dish pre-treated with a gelling agent and incubated at 35°C for 24 hours. Total coliforms per 100 mL of water were calculated by multiplying the resulting number of dark blue, purple, pink and red bacterial colonies by 50. 2.1.7 Macroinvertebrates Aquatic macroinvertebrates are small animals without backbones that are large enough to be seen without a microscope (>1mm) and which live in the mud and gravel at the bottom of waterbodies, within the water column or in the vegetation along the sides of a stream or lake (GLOBE Program, 1997). They are an important part of the aquatic food chain and include the larvae of many insects such as mosquitoes, dragonflies and caddisflies which begin thier lives in the water then become land-dwelling insects when they mature. Other common aquatic macroinvertebrates include crustaceans, snails and worms. Each of these groups of invertebrates have their own environmental requirements and some are very sensitive to changes in pH, DO, temperature, nutrients etc. The presence or absence of these species can therefore be used as an indication of water quality in an aquatic environment. Organisms were sampled according to the type of stream substrate found at a particular site. Sites with a rocky substrate were sampled using the kick sampling method. The sampling site was approached from downstream and a triangular net was placed on the stream bottom. The substrate was disturbed by dislodging stones, etc. with the operator's feet so that any invertebrates attached to the stones were swept by the force of the water into the net. The contents of the net were then transported back to the laboratory. For muddy sites, an artificial substrate consisting of two plastic mesh onion bags, weighted with a brick and enclosed in Nylex™ gutter guard were placed into the stream and left for a period of eight weeks after which the substrate was removed from the stream and transported back to the laboratory. At the laboratory the sample was transferred to white plastic trays and "picked through" for approximately 30 minutes. Macroinvertebrates were identified using Williams (1980) at least to Class level with insects being further classified to the level of Order. Organisms were then counted and ranked according to the Tolerance Ranking Method (Waterwatch Tasmania, 1996) (see Appendix 1). This allowed a biological assessment of water quality at a particular site to be determined. 2.2 VEGETATION SURVEY A vegetation survey was conducted to establish if a correlation existed between vegetation type and the distribution of E. yabbimunna burrows on Shorewell Creek. Because of the relatively small catchment area it was possible to systematically map the riparian vegetation for the entire length of the stream. Vegetation was qualitatively sampled and identified as mosses, ferns, herbs, rushes, grasses, shrubs or trees. These categories were recorded as presence/absence data on to 1:2500 maps produced on a Mapit Geographic Information System (GIS). Areas of homogenous vegetation were then identified and mapped on a 1:10 000 map. Some individual species were identified using Curtis & Morris (1975) and Curtis & Morris (1994). 2.3 BURROW DISTRIBUTION SURVEY In conjunction with the vegetation survey, a systematic search for E. yabbimunna burrows was made in the riparian zone along Shorewell Creek. E. yabbimunna are known to occur in close association with a similar species, E. fosser along Shorewell Creek (Doran & Richards, 1996). Because it was impossible to distinguish between the burrows of E. yabbimunna and E. fosser on the basis of external morphology it was decided to record the location of all burrow sites in the hope that the species responsible could be identified at a later time. At each site where burrows were located a site description and a habitat evaluation were recorded using the data sheets shown in Appendix 2. As the project proceeded it became obvious that it would be desirable to undertake a trapping program in order to establish the presence or otherwise of E. yabbimunna at a particular site. A Scientific Collecting Permit (No. FA 98104), which allowed the excavation of burrows for the purpose of collecting and identifying E. yabbimunna, was issued by the Tasmanian Parks and Wildlife Service under the terms of the Threatened Species Act, 1995. However, given the threatened status of the species, it was felt that a less intrusive capture method would be preferable. It was decided to trial a pitfall trap method used by A.M.M. Richardson (pers. comm.) to capture burrowing crayfish on Tasmania's West Coast. This involved sinking 23 cm plastic flower to the level of the water table at burrow sites. This method had the advantage, that should any crayfish be captured, the water and mud in the bottom of the flowerpot would provide cover from predators and prevent dehydration until the trap could be cleared. As Parks and Wildlife had expressed an interest in being able to distinguish different Engaeus spp. on the basis of burrow morphology it was also decided to trial a trap design based on Norrocky (1984) where a 30cm length of 4 cm diameter PVC drain pipe was fitted with a hinged metal flap which was adjusted to enable it to move freely (see Fig. 6). Click to Enlarge Fig. 6: Burrowing Crayfish Trap The trap was set by placing the pipe into a burrow entrance and covering it with a tin can. The idea was that as the crayfish left its burrow it would push past the metal flap which would subsequently drop back into place behind the animal resulting in the crayfish being trapped in the upper section of pipe. The trapping program took place over a three week period beginning 7/9/98 in areas of observed recent burrowing activity. All traps were set in the late afternoon and checked the following morning.