Diversity of Life in the Litter, Soils, and other Habitats Name Introduction The major organism groups include protozoans, nematodes, rotifers, tardigrades, and the microarthropods. Microarthropods represent a diverse group of organisms, which include five groups that often make a substantial contribution to total community biomass (Wallwork 1976). These five groups are arachnida, myriapoda, apterygote insecta, pterygote insecta, and crustacea. Water Film Organisms Protozoans In terrestrial ecosystems, protozoa are the most common water film organisms and are characteristically associated with the surface layers of litter and soils, in particular decomposing plant material (Lousier and Bamforth 1990). For ecological reasons, we have arranged the protozoans into three groups: flagellates, amoebae, and ciliates. The community structure and pattern of protozoan presence are closely related to ecological conditions of the habitat (Louiser and Bamforth 1990). Protozoans may be the principal consumers of bacteria in the soil, this consumption can help regulate and modify the bacterial community (Stout 1980; Bryant et al.1982; Clarholm 1985; Frey et al.1985; Louiser and Bamforth 1990). Flagellates, naked amoebae, and small ciliates appear to be the most widespread forms and have the largest biomass and highest turnover rates, thus exerting greater impact on the organic cycle of soil (Louiser and Bamforth 1990). Protozoans are able to survive inhospitable periods by forming cysts until conditions become more favorable. Rotifers Rotifers are commonly found in moss and lichen habitats (Gerson and Seaward 1977). Rotifers as a group are generally associated with freshwater but also occur in terrestrial habitats. They are suspension feeders, ingesting bacteria and particles of organic debris (Wallwork 1976). Platt (1968) found that rotifers could feed on lichen ascospores. Gerson and Seaward (1977) estimated that rotifers account for 8.5% of aquatic lichen microfauna. Rotifers are also able to enter a cryptobiotic state when the environment becomes inhospitable.
Tardigrades Tardigrades are micrometazoans found in marine, freshwater, and terrestrial habitats, often called water bears. Even terrestrial tardigrades are essentially aquatic and must be enveloped by a film of water to remain active. Tardigrades feed on soil, mosses, lichens, liverworts, and other vegetation (Nelson and Higgens 1990; Kinchin 1992a, 1992b, 1994). Some large species of tardigrades are predaceous, feeding on protozoans, nematodes, and other tardigrades (Nelson and Higgens 1990). In addition to moss and lichen habitats, tardigrades are common in soil and litter as well. Tardigrades are not evenly distributed in soils, but they are generally found in dense aggregations in one area (Kinchin 1994). When the environment becomes unsuitable tardigrades, enter an anhydrobiotic or more commonly called a cryptobiotic state, known as a tun. Cryptobiosis is an adaptation to enhance survival by allowing the tardigrades to increase longevity and only become active during optimum periods (Nelson and Higgens 1990; Kinchin 1989, 1990, 1992a, 1992b, 1994). Nematodes Nematodes, or roundworms, are generally the most numerous of the larger water film organisms. They are ubiquitous organisms occurring in soil, freshwater, and marine habitats. Nematodes have evolved to fill a wide variety of ecological niches, from free living to parasitic. Terrestrial nematodes feed on a wide variety of foods and can be ecologically grouped as to their stylet and esophageal morphology (Banage 1963; Wasilewaska 1971; Yeates 1979, 1993). Bacterial feeding nematodes, or microbivorous, feed by drawing bacterial suspensions into the intestinal tract using a sucking action. The stylets in this functional group tend to be reduced. Plant and fungal feeders, phytophagous and fungivorous feeders, use a pronounced stylet to pierce the cell wall and suck the contents through the hollow stylet. Omnivorous and predaceous nematodes often have a stylet, teeth, denticles, or some combination of these. They feed on protozoans, tardigrades, rotifers, small oligocheates, or other nematodes (for a review see Freckman and Baldwin 1990). Like tardigrades and rotifers, nematodes enter a cryptobiotic state when environmental conditions become unfavorable. Terrestrial Dry Community Microarthropods The microarthropods account from may of the functional groups that are present within lichen, litter, and soil communities. Of the various subclasses in the arachnida, only three groups play important roles in these microhabitats, the Aranea, Pseudoscorpionida, and the Acari. Both the Aranedia (true spiders) and Pseudoscorpiones (false scorpions) are strictly predators, feeding on smaller soil and litter fauna. They are often the top predators in litter and soil ecosystems (Wallwork 1976; Dondale 1990; Munchmore 1990).
Acari: Mites By far the most important and diverse of this group are the Acari, or the mites. Mites are further subdivided taxonomically and functionally. There are four main groups: prostigmatid mites, mesostigmatid mites, cryptostigmatid mites, and astigmatid mites. The cryptostigmatid mites, called oribatid mites, by far make up the largest of these groups. Ecologically crytpstigmatid mites have low metabolic rates. While they may occur in extremely high numbers and consume large quantities of organic matter, little energy is extracted by these mites (Norton 1990). Their contribution, however, is significant in the pre-processing of organic matter for decomposition by bacteria and fungi (Wallwork 1983; Norton 1986). Cryptostigmatid mites fill a variety of ecological roles but in soil and litter communities they are generally saprophages. The mesostigmatid mites are generally predatory mites and functionally feed on collembola, other mites, nematodes, and eggs of these groups (Wallwork 1976; Krantz and Aniscough 1990). Occasionally, mesostigmatid species are saprophages, bacteriophagous, fungivorous and phytophagous. The prostigmatid mites fill a wide variety of ecological roles, which range from phytophagous to parasitic in nature. This is a highly plastic group, and species within the same genus may be highly variable in their feeding habits (Kethley 1990). Myriapoda: Centipedes, Millipedes, Glass Centipedes, Pauropoda There are four groups within the myriapoda all of which have been associated with soil communities (Wallwork 1976). These include the Chilopoda, Diplopoda, Symphyla, and Pauropoda. The Chilopoda are the centipedes. Centipedes are commonly associated with decaying habitats such as rotting logs and leaf litter. They are among the more important microarthropod predators, but not all species are strictly predaceous and some species switch to a diet of leaf litter during winter months (Wallwork 1976; Mundel 1990; Shelley 1999). The Diplopoda, commonly known as millipedes, are detritovores, specifically phytosaprophagous organisms that feed on decaying plant material. Like the centipedes they are often associated with decaying habitats. They are also commonly found in litter and upper soil horizons (Hoffman 1990). A few species of millipedes are carnivorous, having sucking mouthparts (Shelley 1999). Millipedes often play important roles in the chemical and mechanical conversion of litter into soils (Hoffman 1990). The Symphyla or glasshouse centipedes are mainly omnivorous, feeding on the soft tissues of plants or animals and generally feeding on roots. Other symphylans will feed on detritus in the early stages of decomposition sooner than most other soil invertebrates.
Apterygota Insects(without wings): Springtails, bristletails, and silverfish The apterygota group includes thysanura, diplura, protura, and collembola. Representatives of all four groups of apterygote insects have been found to be associated with the soil and litter microhabitats (Wallwork 1976). Collembola or springtails are arguably the most important group of apterygote insects, and collembola are the second most common soil arthropod, only surpassed by cryptostigmatid mites. Functionally, collembola are saprophages, feeding on a wide variety of decomposing material (Wallwork 1976; Christiansen 1990). It is believed that collembola are generally consuming fungi and bacteria living on the decomposing material rather and the material itself (Testerink 1982; Moore et al.1987; Chen et al. 1996). Pterygote(winged insects): Beetles, Ants, and others A wide variety of pterygote insects can be found in soil and litter at some point in their life history. These may include members from diverse groups such as Diptera, Lepidoptera, and Coleoptera. The most important of these are probably the Coleopterans. Two families: the ground beetles, (Family: Carabidae), and the rove beetles, (Family: Staphylinidae), are important predators within soil and litter habitats (Wallwork 1976). Small Staphylinidae may feed on nematodes, mites, and collembola while larger beetles may feed on adult beetles and large insects as well (Newton 1990).
Wet Extraction Objective: Separate nematodes, rotifers, and tardigrades from soil sample so they can be counted and identified. Materials needed: 20 g (at least) pre-weighed soil (in Petri dishes and labeled) De-chlorinated tap water Rack for holding funnels Screens Clamps Glass funnels Filter paper (Kim-wipes) Scissors Rubber tubing 10% formalin solution Hot plate with stir bar 500 ml beaker Scintillation vials Sharpie pen (permanent marker) Pipette Methods: 1. At least 24 hours before the nematodes are to be extracted fill a carboy or flasks with tap water and leave open to dechlorinate. 2. Place funnels on rack and set screen inside. Funnels should have tubing over end already, but if not put it in place. 3. Clamp end of tube and fill funnel with water. 4. Drain a little bit of water by releasing clamp to be sure water has filled entire tube. Make sure water is just touching bottom of screen. 5. Place soil on Kim-wipe and set on top of screen. Make sure water begins to soak into the filter paper and cover with the Petri dish. Repeat for all samples. 6. Let samples sit for 3 days. 7. Heat 10% formalin solution on hot plate, stirring while heating (use stir bar). 8. Label scintillation vials with the sample number. 9. With the vial carefully remove the clamp from the tubing on the appropriate funnel and fill the vial half full with the liquid in the funnel. Repeat for all samples. 10. Fill the remaining half of the scintillation vial with heated formalin. Cap vials. 11. Bring the samples back to the lab for analysis and clean out funnels and screens for next extraction. Schematic of Nematode Extraction set-up Soil sample Kim-wipe filter screen glass funnel tubing clamp
Arthropod Extraction Objective: Separate arthropods from soil sample so they can be counted and identified. Materials needed: 70% alcohol Rack for holding funnels 4 oz. jars Cheesecloth Aluminum foil Scissors Plastic funnels Paper and pencils Light (heat) source Wire screens Methods: 1. Make sure the rack and funnels are clean and free from dirt. Place each funnel on the rack and put a wire screen inside the funnel. 2. Cut cheesecloth into rectangles (9 inches x 9 inches). Make sure there are enough squares for all the soil samples. 3. Write sample number of the soil sample onto a tiny strip of paper with the PENCIL and place in the bottom of jar. 4. Fill jars 3/4 full with 9:1 solution of 70% alcohol solution. 5. Place a large amount of soil (fist sized) onto the cheesecloth square and wrap up. Cloth should cover the sample. 6. Place the soil sample in the funnel and set the corresponding jar of alcohol NEXT to the funnel. 7. Repeat for remaining samples. 8. Wrap the aluminum foil around the top of the funnel and light source and turn on the light source. 9. Place the jar of alcohol under the bottom of the funnel. 10. The jars need to be checked every day or every other day for alcohol evaporation. DO NOT LET THE JARS DRY OUT. 11. After five days, place the lids on the jars and bring them back to the lab for analysis. 12. Save the soils in the cheesecloth for estimating extraction efficiency and clean the rack and funnels so they will be ready for the next extraction. Schematic of Arthropod Extraction set-up Light Soil sample Cheesecloth foil Wire screen Plastic funnel 4 oz jelly jar (arthropod trap)
Index of Diversity Differences in the abundances of species in communities pose two problems for ecologists. First, the total number of species included: sample size, because as more individuals are sampled, the probability countering rare species increases. Thus we cannot compare diversity areas sampled with different intensities merely by counting species. Second, not all species should contribute equally to our estimate of total diversity cause their functional roles in the community vary, to some degree, in proportion to their overall abundance. Ecologists have tackled the second problem by formulating diversity indices in which the contribution of each species is weighted by its abundance. Two such indices are widely used in ecology: Simpson's index and the Shannon-Weaver index. In both cases, we calculate the indices proportions (p i ) of the species (i) in the total sample of individuals. Simpson s index is: D = 1 2 p i For any particular number of species in a sample (S), the value of D from 1 to S. depending on the evenness of species abundances. When five species have equal abundance, each p i is 0.20. Therefore, each p 2 i = 0.04, the sum of the p 2 i is 0.20, and the reciprocal of the sum is 5, the number of the sample. Similar calculations for some hypothetical. Example: 6 Organisms A=15, B=20, C=10, D=5, E=30, and F=70 Total organisms= 150 Proportions wise Squared Proportions A=15/150=0.10 =0.010 B=20/150=0.134 =0.018 C=10/150=0.067 =0.0045 D=5/150= 0.034 =0.0011 E=30/150=0.20 =0.04 F==70/150=0.467 =0.218 Sum of the proportions is 0.2916 and the reciprocal is 1/0.2916=3.429 So Simpson s Diversity index is 3.429
BIO 101 Diversity of Life in the Litter, Soils, and other Habitats Name: (20 points) 1. Find one organism from your wet extraction mount it on a slide and Carefully Draw and identify to the best of you ability. (Spend some time here stick figures are not acceptable) Magnification Write a brief description of this organism:
2. Find one organism from your dry extraction mount it on a slide and Carefully Draw and identify to the best of you ability. (Spend some time here stick figures are not acceptable) Magnification Write a brief description of this organism: 3. Write a brief description of the micro-habitat you collected your samples from.
4. Organism Counts. As a group count all the organism in both your wet and dry extractions. Outside Class: Using Excel or other program design and make a table you will generate a table to present these results. 5. Calculate the mean number of organism per gram weight 6. Calculate the Simpson s diversity index for your sample.