Cleverson Ranieri dos Santos (MPEG) • E-mail: firstname.lastname@example.org
East Pará - Dr. Cleverson Ranieri dos Santos (MPEG), Dr. Leandro Juen
West Pará - Dr. José Reinaldo Pacheco Peleja (UFOPA)
Amapá - MSc. Inácia Maria Vieira (IEPA)
Maranhão - Dr. Dayani de Fátima Pereira (UFMA)
Groups of interest and diversity of species evaluated per grid: Immature Plecoptera, Trichoptera, Ephemeroptera and Heteroptora (Nepomorpha and Gerromorpha) will be studied, being identified by genera or morphotype. In other Amazon regions, such as the Adolpho Ducke Forest Reserve (RFAD) for example, there are estimated to be around 100 species distributed across various genera among the taxonomic groups mentioned above. Other taxa, such as Odonata, Coleoptera, Heteroptera and Oligochaeta, can be studied in a number of environments in order to understand the substitution of fauna between sampling sites. Freshwater Crustaceans will also be studied, principally the Decapoda; a group which still hasn’t been assessed within the collection grids. Knowledge of crustaceans in the Eastern Amazon is still incipient - only 29 registered species of shrimps and crabs (an introduced species) so far having been dated for the freshwaters of the state of Pará, and only 12 occurrences being registered in the state of Amapá.
Biological role of the group: Plecoptera, Trichoptera and Ephemeroptera are important and abundant components of the aquatic biota, fulfilling an important role in the trophic chain of this particular environment. Furthermore, species from these three orders are among the main groups of macroinvertebrates used in environmental monitoring studies, due to their great sensitivity toward changes in habitat. The aquatic and semi-aquatic Heteroptera are important within ecosystems because of their important role as food for insects, fish, amphibians, reptiles, birds and mammals. They can also play an important biological role in the control of the larvae and pupae of disease transmitting mosquitoes, in the sense that nearly all aquatic species feed on various types of insects.
Invertebrates play a key role within the trophic food chain of freshwater ecosystems, serving as a general determinant in the speed of decomposition of plant remains, as well as being an important source of food for vertebrates. They are also widely used as bioindicators, with various standard responses from different taxa toward environmental changes being known, such as the removal of forest cover and changes to the physico-chemical parameters of water. Some species are important from a health perspective as transmitters of disease, whereas other species are important for controlling these vectors.
Crustaceans are also greatly important within the ecosystem, especially in terms of the food chain throughout their different stages of life. When the crustacean is still larvae or very small (some having direct development) it preys on other small organisms or primary producers, but also serving as food for bigger organisms, acting as a link in the transfer process of energy at various trophic levels. In adult form it can capture invertebrates and even a number of vertebrates, as well as serving as food for various groups of organisms such as man, for example; a number of these species having economic value. The crustacean can inhabit the most diverse of ecosystems, including oceanic waters, estuaries and mangroves, occupying rivers, streams, lakes and even terrestrial environments.
Techniques for collecting aquatic insects
Collection technique 1. Collection of riverbed debris
Samples of riverbed debris will be collected from the stream until a fabric 1mm dipnet (rapiché) can be filled (Figure 1). The riverbed debris collection points sampled during the dipnet (rapiché) filling process must be fully distributed along the transect of the stream, excluding areas of the greatest depth where the dipnet is not able to reach. The surface layer must be sampled, defined here as a section of approximately 10 cm. The number of dipnet samples must be counted along a transect of 50 meters, with the number of samples for every 10 meters being evenly distributed in order to have the same quantity (we suggest 3 full dipnets for every 10 meters). In the event that only small amounts of riverbed debris are found, making it impossible to fill the dipnet, the sampling area must be increased on both sides until a sufficient amount of material can be collected (Figure 3). In the event of it being impossible to fill the dipnet in the sampling area, the required proportion of the obtained sample must be noted (i.e. the filled dipnet percentage) in order to facilitate future evaluations. The collected invertebrates must be preserved in 80% alcohol.
Observation: The portions of debris obtained across different environments must be kept separately in order to study the differences associated with different kinds of environments (different water speeds, depths, shade, surface debris depth, the presence of rocks, types of substrate, etc.). Material will therefore be collected from numerous environments for future study, making sure that the collected quantity corresponds to a full dipnet for each environment in question.
Sampling unit: Collection effort (number of full dipnets) per aquatic plot.
Figure 1. Dipnet design. The netting material at the base must be spaced at 1 mm.
Collection technique 2.
Bottom and surface sediment close to the banks of streams
Samples must be taken from the sediment and/or surface of the water near to the banks of streams, in such situations where environments cannot be satisfactorily sampled using technique 1 - the sampling of riverbed debris over a stream transect. This method consists of a dragging process, as described in the crustacean technique, using 1 mm netting or sieving material. The distance from the bank of the stream must be the minimum possible to facilitate the dragging process. The collected material must be stored in 80% alcohol for laboratory sorting at a later date. It is only necessary to sample the bank of one stream at each collection point when possible substrates are identified (such as debris) containing the target organisms.
Other insects (on the bed or surface) may be sampled in each plot in a non-quantitative manner. The procedure will consist of the straightforward capture of all sighted species, aimed at registering their presence within each plot. The data obtained from these organisms will be treated as present or absent for each plot.
Sampling unit: The dragging effort per 50 meter transect.
Collection technique 3. Collection of adult insects
A small Malaise net (1972 Townes model) will be used, placed at a convenient point along the chosen transect (figure 2). Samples must be removed from each trap every 5 days. The collected specimens will be stored in 80% alcohol until they can be sorted at the laboratory.
Sampling unit: Capture effort per trap every 3 days.
Figura 2. Loaded Malaise trap next to the bank of the stream.
The collections will take place within bodies of water found near to or along the trails within wet regions, but where no stream channel has been positively defined. A 50 m transect will also be used in this particular situation, the collections occurring in the same way as in technique 1, the only difference being that the width of the area to be sampled must be restricted to 2.5 m, measured from the transect line. This width has been stipulated based on the average width of second-order streams. The same method must also be used for Crustaceans.
Sampling unit: An aquatic plot.
Techniques for collecting Crustaceans
Collection technique 4. Bottom dragging with manually operated net (shallow water)
Netting with 2 mm spacing between knots of 3m or 5m will be used (according to the width of the stream in question) at a height of 1.50 m for shallow depths (maximum of 1.50 m). Nets will be operated by two people, one on each side, in contact with the bottom of the body of water for a distance of 10 meters. The net will be immediately closed after use (joining the spreader poles together), then removed from the water. The collected crustaceans will be immediately stored in 80% alcohol and, where possible, pre-sorted into collected taxonomic groups. Sampling should take place within the chosen 50 meter stretch in each plot, using 10 meter subdivisions: 1, 3 and 5 (see figure 4). Assuming it’s not possible to use the type of net described above (depth below 1m or bottom filled with wood or stones), a smaller dragging net must be used (1mm netting), the process then consisting of short 1 to 2 meter dragging actions, with a total of 3 to 5 trawls for each 10 meter stretch, preferably in areas with debris or muddy sediments.
Sampling unit: result of the dragging effort per aquatic plot.
Collection technique 5. Submersed traps
(shrimp nets, gillnets)
This method is for supplementing the stream bed sampling techniques. Submersed traps such as shrimp nets and gillnets, using dead bait (such as fish, for example) will be used. These traps will be placed in subdivisions 2 and 4 of the 50 meter stretches, after the bottom dragging technique has completed (Figure 4). Each trap can be used more than once throughout the day, with the time of submersion being noted on each use, carried out every four hours or, if possible, left overnight.
Sampling unit: the result obtained during the time that each trap remains submersed.
Collection technique 6. Surface sampling
The aim of this technique is to collect small shrimp specimens using an adapted plankton net made out of 200 micron mesh with a 60 cm opening of 1.35 m in length, connected to a flow meter and adjusted for water and surface collection. The net must be launched out, pulled by a 10 meter rope, twice for each plot and within the areas marked out for bottom dragging (Figure 4), making sure there are intervals of at least 30 minutes between draggings.
Sampling unit: the capture effort of each 10 meter sampling area.
Collection technique 7. Manual collection on land
Collections will take place parallel to the plot’s watercourse along a 50 meter stretch at a maximum of 12 meters from the bank of the stream. A collector will walk along one of the corresponding 50 meter stretches, manually collecting crustaceans over dry land by means of a shrimp net. Other routes will be established (up to a maximum of three, if possible), depending on the availability of other collectors, each stretch/collector being separated by three meters, with the first route being closest to the bank (see Figure 4). The time taken by each collector along each route must be standardized, suggesting a total of 10 minutes per 50 meters.
Sampling unit: the result of the efforts of each collector over the traversed distance/time.
Sampling design: A 50 meter area, for aquatic insects and crustaceans alike will be staked out for each stream selected for study, the collections of aquatic macroinvertebrates and environmental parameter measurements (physical and chemical) then going ahead. The plot must start 10 m upstream from the point at which the trail crosses the stream, going in an easterly direction, done so in order to minimize any effects of the trail on results. A 50 m line will be drawn out from this point onwards, with demarcations set out every 10 m, determining the sampling points in accordance with the technical description for each studied group (Figures 2 and 3). For the study of crustaceans, each stretch is subdivided into 5 separate stretches of 10 meters (numbered from 1 to 5, first to last respectively). In the event that the grid has an aquatic ecosystem consisting of pools, lakes or Igapó forest that cannot be subdivided in this way, it will be necessary to define a plot with a measurement system as close as possible to the 50 m system. Between 4 and 10 plots must be established on every site, the overall number being determined, on the one hand, by the complexity of environmental diversity (i.e. sites with various types of environments having larger numbers of plots) and, on the other hand, by the possibility of establishing statistically independent plots (i.e. during periods of drought, for example - the number of possible plots being reduced). There can be a maximum of 10 aquatic plots, each plot distributed as evenly as possible and spread out across the entire grid (figure 5), this suggesting that the hydrography of the area needs to be studied in advance in order to define the plot.
The watercourses chosen for sampling will be those in which the pre-designated techniques can be effectively applied, since deeper watercourses (those greater than 1.60 m in depth) are difficult to sample using the proposed methodology. Sampling independence is achieved by selecting streams from different water basins, or those with the lowest amount of connectivity between them as possible. The geographic coordinates of the sampled stretches will be registered through the use of GPS technology.
Figure 3. Distribution of samples for aquatic insects within each aquatic plot, each of the plots being 50 m in length.
By following the riverbank, six reference points will be set out at a distance of 10 m each. The substrate will then be collected by crossing the watercourse from each of these six points, until the dipnet is filled. The “x” points shown in figure 3 show the points at which the riverbed debris can be collected, with a maximum 3 points every 10 meters. The 20 m and 30 m reference points are indicative of a watercourse that is deeper than the reach attainable by the dipnet, illustrated in figure 3 by a stretch where no substrate collection can be made.
Figura 4. Collection technique usage model for crustaceans in a single plot, comprising of a sampled watercourse of 50 meters in length, separated into 5 subdivisions (1 to 5) of 10 meters each
Three water collection techniques will be used, two of these (bottom dragging and surface collection) within subdivisions 1, 3 and 5; it being necessary for the second applied technique to utilize a time gap of at least 3 hours after application of the first. Submersed ‘waiting’ traps will be place at subdivisions 2 and 4, with a minimum distance of 3 meters between each one. The land collection technique will follow a line along one of the banks at a distance of up to 12 meters from the water itself, in addition to a gap of approximately 3 meters between each route taken by the collectors.
Figura 5. Hypothetical layout of the plots chosen for sampling, showing four sampling points in grid (A) and eight in grid (B). The layout will vary depending on the hydrographic area in question.
Additional and important environmental data for this group: Environmental data from each plot will be collected in order to carry out a minimum characterization study of the sampled areas. The average width of the channel (m) will be calculated from the average value of 3 equidistant measurements along the stretch in question. The average depth of the channel (m) as well as the average maximum depth (m) will be calculated by means of 4 equidistant surveys at 3 transversal transects, also equidistant, along the stretch in question. The speed of the current (m/s) will be determined by averaging three points set out within the longitudinal center of the channel and measured in the middle of the water column, done so through the use of a flowmeter or by measuring the travel time of an object floating over a known distance.
The average flow (m3/s) will be obtained from a combination of average speed, width and depth, calculated using the formula: Q = A . As; in which Q = flow; As = Average speed of the current; A = average trans-sectional area within the watercourse cross-section. The trans-sectional area will be calculated from the average area of 3 transects in each studied stretch, by the formula: At = Sn iAn, in which:
At = transect area, given by the sum [(Z1+Z2)/2].l + [(Z2+Z3)/2].l + … [(Zn+Zn+1)/2].l, in which:
Zn = depth measured in each segment and l = the width of each segment.
The type of substrate will initially be classified into seven categories: sand, clay, trunks (pieces of wood with diameter greater than 10 cm), litter (consisting of leaves and small branches), fine litter (fine particulate material), roots (tangled roots, normally fine, deriving from riverbank vegetation) and macrophytes (aquatic vegetation). The following variables will also be obtained: the potential hydrogen (pH), conductivity (μS/cm), dissolved oxygen (mg/L) and background and surface temperatures (°C), being determined using specific types of portable equipment. Other types of environmental characteristics may also be included such as wind strength and direction, weather conditions and water turbidity, for example.
Method of preserving the collected material: In the field, the collected material will be preserved in wet medium of 80% or 90% ethanol, later preserved in 80% ethanol at the laboratory after the insects have been sorted and identified. After initially being stored in 90% alcohol, the crustaceans will then be removed, identified and stored in 80% alcohol. The collected samples will then be deposited in MPEG and other trustworthy Amazon depository collections.
Restrictions on activities that could prejudice protocol development: Other teams carrying out activities in the area, making sure not to set foot in the streams that are part of the stretches or any part of the stretches that are yet to be sampled.
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MELO, G. A. S. (Ed.). Manual de Identificação dos Crustacea Decapoda de água doce do Brasil. São Paulo: Loyola, 430 p.
MORIN, A.; STEPHENSON, J.; STRIKE, J.; SOLIMINI, A.G. Sieve retention probabilities of stream benthic invertebrates. J. N. Am. Benthol. Soc., v. 23, n. 2, p.383-391, 2004.
TOWNES, H. A light-weight Malaise trap. Entomology News, v. 83, p. 239-247, 1972.