A STUDY OF THE DISTRIBUTION OF NAEGLERIA SP. IN SOIL BY Ndu Umeche A.DISSERTATION Submitted to Michigan State University in partial fulfillment of the requirements for the degree of DOCTOR OF PHILOSOPHY Department of Zoology 1978 ABSTRACT A STUDY OF THE DISTRIBUTION OF NAEGLERIA SP. IN SOIL By Ndu Umeche A study was conducted at Rose Lake Wildlife Research Area in Michigan to ascertain the distribution of Naegleria sp. in different types of organic and mineral soils, to estimate their populations and to characterize some factors influencing their distribution in soils. Core samples taken throughout the year did not reveal the presence of high temperature strains of g, gruberi or pathogenic strains of N, figulggi. Four non-pathogenic fl, gruberi isolates were obtained: 3 from mineral soils (Ottokee loamy sand, Fox loam and Miami loam) and 1 from organic soil (Rifle peat). Percentage moisture, pH, bulk density and porosity of soils were determined. The isolates were obtained from sites with litter and soil layers containing 17 and 35% moisture levels, whereas other sites with higher moisture levels up to 177% contained predominantly Colpoda sp. and other protozoa. The number of Naegleria and Naegleria- like amoebae (amoebae that resemble Naegleria in cyst and tropho- zoite morphologies but did not transform into the flagellate stage) per gram of litter and soil were estimated using a dilu- tion method by Singh (1946). The populations which ranged from Ndu Umeche 159 to 16,000 per gm varied from soil to soil, layer to layer and time to time, with more cells in the soil than in the litter layers. Using autoclaved soils, inoculation of eria isolates per gm of organic and mineral soils in excess of population range in nature with bacterial food added resulted in death of cells and lack of growth. Inoculation of cells within natural population range with bacterial food added resulted in growth of cells, although the increase in number was not significant. These results suggest limited growth of the amoebae in soil environments, whereas they grew rapidly in pure laboratory cultures. Using unautoclaved soils, inoculation of Naegleria isolates per gm of organic and mineral soils with no bacterial food added resulted in significant growth in samples from positive sites and poor growth or death of cells in samples from negative sites. An inference was made that the positive sites might contain edible bacteria or other edible materials that favored growth, while negative sites might contain inedible bacteria or other toxic materials that inhibited Naegleria development at those sites. ACKNOWLEDGEMENTS I wish to express my appreciation and thanks to my major advisor, Dr. R. Neal Band for his guidance and helpful suggestions in the course of this research. Constructive criticisms by Dr. J. W. Butcher, Dr. T. W. Porter and Dr. J. A. Breznak were very helpful in the preparation of this thesis. My thanks are extended to Dr. Pedro Galindo and Dr. Carl JOhnson of Gorgas Memorial Laboratory, Canal Zone, Panama, for providing their research facilities and assistance to conduct a part of this study. Dr. E. P. Whiteside assisted in the analysis of Michigan soils. Finally, I wish to thank the Department of Zoology, Michigan State University, for financial aid and members of my family for their moral support. ii TABLE OF CONTENTS LIST OF TABLES . . . . . . . . . . . . . . . . . . . . LIST OF FIGURES . . . . . . . . . . . . . . . . . . . INTRODUCTION . . . . . . . . . . . . . . . . . . . . MATERIALS AND METHODS . . . . . . . . . . . . . . . . Sampling sites . . . . . . . . . . . . . . . . . Sampling methods . . . . . . . . . . . . . . . . Culture methods . . . . . . . . . . . . . . . . . Culture media . . . . . . . . . . . . . . . . Soil sample preparation . . . . . . . . . . . ' eria isolation . . . . . . . . . . . . . Protozoa count . . . . . . . . . . . . . . . . Growth in autoclaved and unautoclaved soils . Hemacytometer count and dilution method count Percentage moisture determination . . . . . . . . pH determination . . . . . . . . . . . . . . . . Bulk density and porosity determination . . . . . RESULTS . . . . . . . . . . . . . . . . . . . . . . . Sampling sites and soil types . . . . . . . . . . Distribution of life by strata . . . . . . . . . Characteristics of the four Nggglggia isolates . Vegetation cover . . . . . . . . . . . . . . . . iii Page vii 10 10 10 ll 11 12 12 13 13 13 15 15 15 16 17 Percentage moisture . . . . . Bulk density and porosity . . pH determination . . . . . . Naegleria growth in the soil Numbers per gm dry soil . Growth in autoclaved soils Growth in unautoclaved soils D ISCUSS ION O O O O O O O O O O O Nae Naegleria species per gm of soil e ia distribution and isolation Physichochemical factors . . Numbers per gm soil . . . . Growth in autoclaved soils . Growth in unautoclaved soils SUMMARY LIST OF REFERENCES . . . . . . . . APPENDICES A. B. A table for estimating protozoal numbers Actual protozoa count . . . . iv Page l7 18 18 18 18 19 19 54 54 56 57 58 59 59 61 63 69 71 10. 11. 12. 13. LIST OF TABLES Soil samples and soil descriptions . . . . . . Distribution of life in the soils by strata (sept-DEC) 1976 o o o o o o o o o o o o o o 0 Distribution of life in the soils by strata (Jan-Mar) 1977 o o o o o o o o o o o o o o 0 Distribution of life in the soils by strata (Apr-Jun) 1977 o o O o o o o o o c o o o o 0 Distribution of life in the soils by strata (JUl-Aug) 1977 o o o o o o o o o o o o o o 0 Characteristics of the four Naegleria isolates Characteristics of the vegetation cover of organic and mineral soils (Nov. 1977) . . . . Percentage moisture in different strata of organic and mineral soils (Oct. 1977). . . . . Comparison of percentage moisture fluctuation in organic and mineral soils (Oct/Nov, 1977) . Bulk density and porosity of organic and mine- 1'31 80118 (NOV. 1977) o o o o o o o a o o o 0 pH of selected organic and mineral soils (NOV. 1977) o o o o o o o o o o o o o o o o c Number of Naegleria and Naegleria-like cysts and vegetatives corrected to number per gm dry soil in different layers of positive samples. E, 991} added. (Aug/Sept 1977) . . Estimation of growth of inoculated Naegleria isolates into different layers of autoclaved organic and mineral soils. Cells inoculated in excess of population range in nature. ' E: ggli_added. (Oct. 1977) . . . . . . . . . . Page 21 23 24 25 26 27 28 29 30 31 32 33 LIST OF TABLES (cont'd) Table 14. 15. 16. Page Estimation of growth of inoculated Naegleria isolates into different layers of autoclaved organic and mineral soils. Cells inoculated within population range in nature. E, 92;; added. (Oct. 1977). . . . . . . . . . . . . . . . 35 Estimation of growth of inoculated Naegleria isolates into different layers of unautoclaved organic and mineral soils. No bacterial food added . . . . . . . . . . . . . . . . . . . . . . 36 Comparison of hemacytometer count and dilution methOd count (oct. 1977) I O O C O O I O O O C O O 37 vi Figure LIST OF FIGURES l. A.map of Rose Lake Wildlife Research Area with numbers indicating sampling sites .. . . . . . 2. ll. 12. 13. 14. 15. 16. A photograph of site (Mineral soil) . . . A profile of site #7 (Mineral soil) . . . A photograph of site 8011 O O 0 O O O O O A profile of site #6 A photograph of site A profile of site #8 A photograph of site #10 Miami loam (Mineral soil) #7 Ottokee loamy sand Ottokee loamy sand #6 Carlisle muck (Organic Carlisle muck (Organic soil) #8 Rifle peat (Organic soil) Rifle peat (Organic soil) . . A profile of site #10 Miami loam (Mineral soil) A photograph of site CMineral soil) . . . #13 Hillsdale sandy loam O O O O O O O O O O O O O A profile of site #13 Hillsdale sandy loam (Mineral soil) . . . A photograph of soil 8011 O C O O O O O . #15 Sebewa loam (Mineral A profile of site #15 Sebewa loam (Mineral 8011 O O O O O C O O A diagram of amoeba cell of Naegleria gruberi A diagram of flagellate cell of.fl, gruberi . . A diagram of cyst of N5 gruberi . . . . . . . vii Page 39 41 41 43 43 45 45 47 47 49 49 51 51 53 53 53 INTRODUCTION Protozoa have been isolated and described from soil cultures since the later part of the nineteenth century (Muller, 1887). In experiments with partially sterilized soils, protozoa were found to be one of the factors limiting the development of bacteria in soils and this limita- tion affected soil fertility (Russell and Hutchinson, 1909). Most protozoa cultured from soils developed from resting cysts (Goodey, 1911). Examination of freshly fixed soil films showed that protozoa could also exist in an active trophic state in normal soils (Martin and Lewin, 1915). Although a few protozoa in soil samples could be observed directly under the microscope by staining (Heal, 1964), most of them were observed by incubating soil samples in various media at various tempe- ratures for several days. Nutrient agar, soil extract agar and hay infusion liquid media were commonly used depending on the organism. In some studies, the soil samples were incubated on the media and the protozoa obtained their food from the indigenous bacterial flora of the soil (Cutler, 1919; Crump, 1920). In other studies, the soil samples were incubated with pre-grown bacterial cultures (Cunningham, 1915; Cutler, 1923). Some methods had been developed to estimate the protozoa populations in the soil. A.method of enumeration similar to that used in estimating bacterial numbers was described by Cunningham (1915). In this method, bacteria were inoculated into soil extract medium and incu- bated for two days. Then a series of soil dilutions was made in test 2 tubes and inoculated into the pre-grown bacterial cultures and incu- bated for several days before counting the protozoa. To obtain counts of cysts only, the soil samples were heated to 58°C to kill the vege- tative cells. However, it was found that heating also killed a consi- derable number of cysts. A.more reliable method of eliminating the vegetatives without affecting the cysts consisted of treating the soil samples with 1.5 to 2.0% hydrochloric acid for 24 hr. (Cutler, 1920). The dilution method was modified further by Singh (1946) using Fisher and Yates (1943) Statistical Table VIII2 for estimating the densities of microorganisms. Singh's method involved a twofold dilution series of soil samples ranging from 1/5 to l/81,920. Each dilution series was inoculated into 8 glass rings embedded in agar plates containing edible bacteria. After incubation, sterile and fertile rings were counted and the number of protozoa per gram of soil was estimated with the Table from the count of negative cultures. Two estimates would differ significantly at the 5% level when their numbers of negative cultures differed by 8. Excellent estimates of protozoa numbers have been obtained by various workers using the dilution method with appro- priate modifications. Soil protozoa feed mainly holozoically, although saprozoic nutri- tion occurs in some species. Holozoic feeders like amoebae and ciliates ingest bacteria and other small organic materials, while saprozoic feeders like some flagellates obtain their nutrients by absorption. Protozoa play a dominant role in regulating the bacterial population in the soil. This was confirmed by studies in which known numbers of protozoa and bacteria were inoculated into sterilized soils: as the -3 numbers of protozoa increased, the bacterial numbers decreased; in soils without protozoa, a high bacterial density was maintained (Cutler, 1923; Danso and Alexander, 1975; Habte and Alexander, 1977). Some bacteria are eaten by soil protozoa, but others are not. Both Gram-positive and Gram-negative bacteria are eaten (Singh, 1941). Many pigmented species like Chromobacterium violaceum and toxin-producing species like Serratia marcescens are not eaten. Pseudomonas pyocyanea produces pyocyanin pigment and other metabolic products that are toxic to soil amoebae, flagellates and ciliates (Singh, 1942; Singh, 1945). When food is lacking, encystment or death takes place; in the presence of edible bacteria, bacterial extracts or amino acids, excystation occurs and growth takes place (Singh g£_§1,, 1971). Studies of the rhizosphere of plants using the buried slide tech- nique revealed the presence of large numbers of cysts and vegetative protozoa (Starkey, 1938). The protozoa numbers and diversity were higher in the rhizosphere than in the surrounding soil. The most abun- dant populations occurred around decaying roots where rapid bacterial multiplication was favored, although the abundance of particular genera varied with plant species and age (Geltzer, 1963; Darbyshire and Greaves, 1967). The protozoa were probably attracted to the rhizosphere by the abundant bacterial population there or as a consequence of the nutrients diffusing out of the root cells, or both (Linford, 1942). Most protozoa are found between the upper one inch and twelve inches of soil. This region contains most of the organic materials and nutrients that favor development of bacteria on which protozoa subsist. It is not unusual to find practically no protozoa below twelve inches 4 in some orchard and garden soils (Waksman, 1916). Protozoa have also been obtained from excavation depths of over twenty feet in ranch soils, but their maximum numbers occurred at a depth of four inches (Kofoid, 1915). A study of protozoa populations in a shortgrass prairie using core samples indicated that the numbers of cysts did not differ with depth, but the numbers of vegetatives decreased significantly with depth (Elliott and Coleman, 1977). The presence of protozoa increased the amount of atmospheric nitro- gen fixed by Azotobacter chrococcum and.A. vinelandii in soil cultures. Cultivation of various species of protozoa with Azotobacter in artifi- cial culture media or in sand cultures and estimation of the nitrogen by Kjeldahl method indicated fixation of 361 or more over control plots. The factors inducing these large fixations were not completely known, but it was believed that a symbiotic relationship was involved between the protozoa and the bacteria. The protozoa might be reducing the acidity of the media and such reduction would lead to increased growth and fixation by Azotobacter. The protozoa might also be removing some of the metabolic products or wastes formed by Azotobacter thereby increasing their nitrogen-fixing efficiency (Nasir, 1923; Cutler and Bal, 1926; Hirai and Hino, 1928). Soil is a complex environment consisting of inorganic (mineral) and organic components. Soils with predominantly mineral constituents and about 1 to 10% of organic matter are called mineral soils. The mineral particles, namely clay, silt and sand are bound into aggregates by organic matter, adsorbed cations and slimy surfaces of microorganisms. The structure of the soil is determined by the size, shape and 5 arrangement of the aggregates. The aggregates also determine the amount of pore spaces available for moisture and aeration (Griffiths, 1965). Soils that contain a high amount of organic matter, about 80 to 951 are referred to as organic soils. The two major types are peat and muck. Peat consists of partially decomposed identifiable organic matter, while muck consists of organic matter in advanced stages of decomposition such that the organic matter could not be identified (Dawson, 1956). The principal sources of organic matter are fallen leaves, animal remains and excreta. Organic matter is the major site of activities of soil microorganisms (Robinson, 1949). The activities of protozoa in the soil are influenced by some physi- cal and chemical factors. Moisture is considered to be the most impor- tant factor; protozoan populations increase significantly when water is added, and die or encyst during periods of water stress (Band and Umeche, 1976; Elliott and Coleman, 1977). The moisture holding capacity of organic soils is about two times or more greater than that of mineral soils (Feustel and Byers, 1936). Since protozoa are aerobic, oxygen from the atmosphere diffuses through the soil and is used for metabolism, while carbon dioxide diffuses out in a similar way (Griffin, 1963). ‘Most protozoa are found in environments around pH 6 to 8, although a few species survive at lower or higher pH values. The temperature of the soil varies from season to season, but greater fluctuations occur at the upper layers than at the deeper layers (Buckman and Brady, 1969). High tempe- ratures tend to increase the metabolic rate while low temperatures slow it down, for example,.gglpgda inflata multiplied three times faster at 27°C than at 10°C (Lackey, 1938). 6 Four major types of protozoa are found in soils: the flagellates, the ciliates, the testate amoebae and the naked amoebae. The flage- llates possess one or more flagella and swim or creep in moisture films. They feed holozoically or saprophytically. The common soil genera are Cercomonas, Oikomonas and Heteromita. All form resistant cysts in adverse environments. Some species such as Heteromita globosa form resting structures that resemble cysts by losing the flagella and rounding up. These structures are less refractile and less resistant than cysts (Sandon, 1927). Flagellates number from 100 to 10,000 per gm in field soils (Crump, 1920). Colpoda is the most common and most predominant of all ciliate genera. Other genera reported in the literature are Blepharisma, Metopus, Halteria and Dileptus. They feed mainly on bacteria, but some feed on amoebae, algae and smaller ciliates (Bick and Buitkamp, 1976). Species of Colpoda are widespread in many types of soil because they can tolerate a wide range of environmental conditions and also form cysts. Examples of Colpoda resistance were shown by studies in which dry soil and hay samples stored for several decades contained viable Colpoda cysts (Goodey, 1915; Dawson and Hewitt, 1931). Some ciliate species such as Blepharisma are more specialized in their environmental requirements, for example, they prefer places with low oxygen tension (Stout, 1958; Noland, 1925). Ciliates number from 100 to 5,000 per gm in forest soils (Bamforth, 1971). Testate amoebae are amoebae that possess shells or tests. The common soil genera are Euglypha, Difflugia, Arcella, Assulina, 322213 and Centronxi . They construct their tests with silica, sand and 7 other organic and mineral particles in the soil. Availability of test materials influence their distribution in soils. Stump (1936) showed that in cultures lacking test materials, Pontigulasia.ya§ failed to reproduce, but when test materials such as powdered sand and glass were added, reproduction started to occur. Extensive surveys of soil testacea have been made by Heal (1961), Heal (1962), Heal (1964) and Heal (1965). Their habitats ranged from bog areas (pH 3.2 to 4.6) to marble soil (pH 6.0 to 8.5). Their classification is based on test morphology, for example, Centropyxis and Plagiopyxis have flattened tests while Difflugia and.Ngbgl§ have pyriform tests. Mbst testaceans respond to unfavorable conditions by encysting. Encysted forms are rounded with closed tests. These forms could be distinguished by staining the animal with phenolic aniline blue on agar film slides. Total numbers of testacea recorded from grassland soils ranged from 40,000 to 69,000 per gm dry soil, while woodland soils ranged from 4,000 to 31,000 per gm dry soil. Naked amoebae are the most numerous protozoa in the soil, where they feed mainly on bacteria. Common soil genera are Naegleria, Hartmanella, Vahlkggpfia, Acanthamoeba and some unidentified'égggbg sp. Some genera such as Naegleria possess a flagellate stage and are called amoeba-flagellates, but most lack a flagellate stage. The naked amoebae are considered to be the most important of all soil protozoa because they play a major role in controlling bacterial population in the soil. Their number may reach as high as 230,000 per gm in soil (Cutler, 1923; Sandon, 1927). Naegleria gruberi is an amoeba-flagellate which has been described 8 as one of the most common soil amoebae. It was first isolated from diarrhoeic faeces by Schardinger (1889). Since then, various strains have been isolated from the soil and other environments world-wide (Rafalko, 1947; Singh, 1952; Chang, 1958; Schuster, 1963a, b; Kingston and Warhurst, 1969). A pathogenic strain of Naegleria designated as .N. fowleri was isolated recently from the soil by Anderson and Jamieson (1972). From soils, they contaminate freshwater pools and lakes where humans swim and become infected. .N. fowleri cysts and vegetative cells were recovered from the cerebrospinal fluid of patients who died from the amoebic meningoencephalitis (Culbertson.g§_§1., 1968; Butt‘g§_gl., 1968; Callicott g§,a ., 1968; Carter, 1968; Cerva and Novak, 1968; Symmers, 1969; Dos Santos, 1970). .N. gruberi has a triphasic life cycle: the amoeboid, the flagellate and cystic phases. The vegetative organism ranges in size from 8 u to 35 u. The cyst has a double wall with one or more pores and ranges in size from 8 u to 16 u. The amoeba moves with broad pseudopodia, feeds by engulfing bacteria and reproduces by fission. The flagellate is a non-feeding stage which lasts from one to several hours. Transforma- tion from the amoeboid to the flagellate stage occurs when the bacterial nutrients are washed off and the amoebae are suspended in distilled water or buffer solutions at room temperature. The flagellate stage may change back to the amoeboid stage or may encyst (Rafalko, 1947). The objectives of the present study were: 1) to isolate pathogenic and non-pathogenic strains of Naegleria from different types of soil, 2) to enumerate Naegleria populations in these soils and 3) to characte- rize some factors influencing Naegleria distribution in these soils. MATERIALS AND METHODS SAMPLING SITES: All soil samples were obtained from Rose Lake Wildlife Research Area. The Area consists of 3,334 acres of forests, grasslands, marshes and lakes located 12 miles northeast of Lansing, Michigan. Twelve sampling sites were selected and identified with the aid of a Michigan Department of Natural Resources (1969) map of Rose Lake Wildlife Research Area, Johnsgard gt a1., (1942) Clinton County Soil Survey, and Threlkeld and Feenstra (1974) Shiawassee County Soil Survey. The sites spread out over the entire area and are interspersed by hard-surfaced roads, gravel roads and trails which are easily acce- ssible. Tags were placed on trees at sites for easy identification. Samples were taken monthly throughout the year. SAMPLING METHODS: Soil samples were taken with modified core samplers made of metal pipes 18" long and 2" diameter. An opening 1/2" wide was cut in each pipe for observation of the profile. To take a sample, the vegetation cover and litter were carefully removed, the sampler was held vertically and driven with a hammer into the ground down to a depth of 12". It was then pulled out gently, wrapped with poly- ethylene sheets and placed into a bucket. The litter was collected and placed on top of the profile or in separate container. The vegetation cover was identified using methods described by Symonds (1958). Samples were transported to the laboratory where they were analyzed the same day or stored in a cold room at 8°C for later analysis. During the months of January, February and March, 1977, deeper layers of the soils were frozen; 10 only the litter and the 1" soil layers were sampled. CULTURE METHODS: Culture media: All soil samples and the protozoa isolates were incubated at 23°C and 37°C on Dilute Stock Agar Glucose - DSAG (Balamuth, W. Personal Communication) which was composed of MgC1.6H20 2.13 g, KH2P04 0.136 g, NaZHP04 0.568 g, trypticase l g, yeast extract 1 3, glucose 1 g, agar 15 g, and distilled water 1L. The DSAG was enriched with Escherichia coli K12 suspension in Low Salt - L.S. (Band and Mohrlok, 1969) composed of NaCl 2.92 g, Mg804 0.65 g, CaC12 0.04 g, and distilled water IL. The DSAG was autoclaved in a flask and poured into sterilized culture plates. E, 92;; was grown on sterilized Stock culture agar (Difco) slants in test tubes at 37°C for 24 hr. The bacterial cells were harvested with a loop, placed into L.S. and shaken with a Vortex mixer. Soil Sample Preparation: Soil samples were prepared for incubation by the following modification of the dilution method described by Singh (1946): ‘warm'DSAG was pipetted into wells of Falcon Multiwell Tissue Culture plates just enough to cover the bottoms. When the agar solidi- fied, the wells were labelled in groups of 8, since 8 replicates of each dilution series would be made. Then 0.3 ml of E, coli suspension was added into each well. Some soil samples were taken out through the openings in the core samplers at the l" and the 8" markings and placed into watchglasses. Big rocks and stones were removed from the samples. One gram of each soil sample was weighed out on a balance and placed into sterilized test tubes containing 5 ml of L.S. The soil in L.S. was mixed on a Vortex ll mixer and then centrifuged. Each litter sample was immersed in L.S. on a 1:5 ratio and mixed by grinding inside a Waring blendor. Five ml of the ground litter was placed into a test tube. Fifteen, twofold dilution series were made from the soil and litter 1:5 dilution, i.e. series ranging from 1/5 to 1/81,920. Then 0.05 ml of each homogenate was pipetted out and added into each of the 8 appro- priately labelled Falcon wells containing E; 9211 suspension. The plates were shaken slightly and incubated at 23°C and 37°C for 48 hr. After incubation, any excess fluid was drained off and the plates were examined directly under a phase contrast microscope. Naegleria and Naegleria-like cysts were transferred to glass slides and identified under high power. Naegleria Isolation: To isolate Naegleria, amoebae or cysts were transferred with a loop from positive Falcon wells to DSAG Petri dishes. A few drops of E. 221; suspension were spread on the cells and the dishes were incubated for 24 hr. Serial subculturing continued on fresh DSAG Petri dishes until pure isolates were obtained. The amoebae were washed three times in L.S. by centrifugation and transformed into flage- llates in a rotary shaker maintained at 28°C. Stock cultures of all isolates were stored on.DSAG Petri dishes inside a refrigerator at 8°C and subcultured monthly. Protozoa Count: The number of Naggleria and Naegleria-like amoebae present in l g of soil or litter from positive sites was determined by application of Singh's (1946) method, and Fisher and Yates (1943) Statistical Table VIII2 for estimating the densities of microorganisms. 12 Positive and negative wells of the 8 replicate cultures per dilution series were counted. Total numbers of both cysts and vegetatives were determined and corrected to number per gm dry soil. To obtain cysts only, each sample was treated with 1% solution of sodium dodecyl sul- fate (SDS) in order to kill the vegetative cells and leave the cysts unharmed. Treated samples were washed three times with L.S. by centrifugation and serially diluted as described above. Growth in Autoclaved and Unautoclaved Soils: Soil samples were autoclaved for 3 hr. at 121°C and 15 lbs/sq. in. pressure. Naegleria isolates from #8 Peat (Org) and #7 Loamy sand (Min) were grown, washed and counted by using a hemacytometer. Various numbers of cells of #8 Peat (Org) isolate were inoculated into each gram of autoclaved and unautoclaved organic soils from different layers, while cells of #7 Loamy sand (Min) isolate were similarly inoculated into mineral soils. The bacterial food was added to the autoclaved soils but none was added to the unautoclaved soils. The inoculated samples were incubated at 23°C for 48 hr. and then serially diluted as described. Hemacytometer Count and Dilution Method Count: To test the relia- bility of the counts obtained by the dilution method, a "recovery test" was conducted. This consisted of counting cells of the two isolates on a hemacytometer and inoculating known numbers of the cells into each gram of autoclaved soils in order to see how many cells that would be recovered. The inoculated soils were serially diluted immediately and incubated. After incubation, the actual numbers of negative cultures observed were compared with the numbers of negative cultures that would be obtained by calculations. The result outlined in Table 16 shows the 13 hemacytometer count inoculated, the expected and the observed negative cultures by the dilution method count, and the differences in negative cultures. It can be seen that the differences in negative cultures were significant in only two samples: #8 Rifle Peat (Org) at l" and #7 Ottokee Loamy Sand (Min) at 8" layers. The remaining six samples did not differ significantly. Two estimates would differ significantly at the 5% level when their numbers of negative cultures differed by 8. Since the differences were not significant in most instances, it was concluded that reliable estimates of the amoeba numbers could be obtained with the soil samples. PERCENTAGE MOISTURE DETERMINATIONS: Soil and litter samples obtained as described earlier were transported to the laboratory in metal containers to prevent moisture loss. Each wet sample was weighed and placed into a pre-weighed watchglass. The weighed samples were dried for 24 hr. inside an oven set at 110°C. After drying, the weights of dried samples were determined and subtracted from the weights of wet samples. The percentage moisture was then calculated. pH.DETERMINATION: Soil samples taken out from the 1" and 8" layers of the core were mixed thoroughly on a 1:1 ratio with double distilled water in small beakers. Litter samples were mixed by a similar ratio but ground up in a Waring blendor. The pH of each homogenate was read from a pH meter. BULK.DENSITY AND POROSITY DETERMINATION: Bulk density and porosity were determined by a modification of the Keen and Raczkowski (1921) box method. Cardboard boxes were used instead of brass boxes. Each box was made as follows: A square 8 cm x 8 cm was cut out from the cardboard l4 sheet. Four rectangular pieces 8 cm x 4 cm were also cut out and glued to all sides of the square with a masking tape. The volume of the box was 256 cc. To compare the volumes of all boxes, each box was filled with water which was then poured into a graduated cylinder and read. All volumes were approximately the same. The boxes were weighed, labelled and taken to the sampling sites. Samples were obtained by cutting out approximately 8 cm x 8 cm x 4 cm pieces of soil with a shovel, lifting and trimming the samples with a spatula and placing them into the boxes. The samples were placed into metal containers with lids for transportation to the laboratory where they were weighed immediately. Small holes were punctured through the boxes at all corners to ensure even distribution of heat throughout the soil. The samples were dried at 110°C for 24 hr. The oven-dried soils were cooled briefly at room temperature and weighed again. The bulk density and porosity were then calculated by methods of Richards (1969). RESULTS SAMPLING SITES AND SOIL TYPES: The twelve sampling sites comp- rised different types of organic soils (peat and muck) and mineral soils (loamy sand, sandy loam and loam). The locations of the sites and detailed descriptions of the soil types are outlined on the map of Rose Lake Wildlife Research Area (Figure 1 and Table l). The photo- graphs of some representative sites and their corresponding profiles are presented in Figures 2 through 13. DISTRIBUTION OF LIFE BY STRATA: The layers of all organic and mineral soils studied throughout the year did not reveal the presence of high temperature strains of N. gruberi, or N} fowleri. However, low temperature, free-living strains of.N, gruberi (Figures 14 through 16), Naegleri -like amoebae and other protozoa were obtained at different times. Naegleri -like amoebae refer to two species of amoebae that resemble N. gruberi in both cyst and trophozoite morpho- logies, but did not transform into the flagellate stage. One of the two species designated as (N-lk) has the same size as Nb gruberi, while the other species designated as (N-lk sml) is smaller than u, gggberi. From September to December 1976, N, gruberi was isolated from only three sites, namely #7 Ottokee loamy sand, #8 Rifle peat and #9 Fox loam; numerous Na r a—like amoebae, Acanthamoeba and the ciliate Colpoda were present at almost all sites in various layers (Table 2) From January to March 1977, N, gruberi was still present at the 15 16 three positive sites and absent at all other sites; all sites showed dominance of Colpoda and Acanthamoeba , while Naegleria-like amoebae were present at some sites (Table 3). From April to June 1977, EL gruberi was isolatable from only one of the three positive sites, namely #7 Ottokee loamy sand; all sites contained predominantly Colpoda; Naegleri -1ike amoebae were present at some sites; the ciliate Blepharisma was seen in the litter layers of #15 Sebewa loam (Table 4) From July to August 1977, N: gruberi was isolated from the #10 Miami loam site; it was also present at all three sites of initial isolation; Naegleri -like amoebae, Acanthamoeba and Colpoda were pre- sent. During this period, Blepharisma was still present at site #15 Sebewa loam (Table 5). Therefore, the months of July to August appeared to be the growing season for all the protozoa observed. CHARCTERISTICS OF THE FOUR NAEGLERIA ISOLATES: Three of the .N: gruberi isolates were obtained from mineral soils (#7 Ottokee loamy sand, #9 Fox loam and #10 Miami loam), while one came from organic soil (#8 Rifle peat). In both mineral and organic soils, Naegleria could be obtained from the 1" and 8" layers at different times. However, in the mineral soil (#10 Miami loam), it was found only in the litter layers (Table 6). The time of transformation from amoebae to flagellates was as expected in all cases. The dilution series from which the isolates originated were as follows: 1/80 for #7 Ottokee loamy sand, 1/5 for #8 Rifle peat, 1/40 for #9 Fox loam and 1/2,560 for #10 Miami loam. l7 VEGETATION COVER: The vegetation cover of sampling sites was composed of grass, leaves or a mixture of grass and leaves (Table 7). The following types of leaves were observed at the indicated sites: White poplar (Soil #5 Houghton muck), Elm (#7 Ottokee loamy sand), Ash (#11 and #14 Bellefontaine sandy loam), Oak (#12 Bellefontaine S.L. and #13 Hillsdale S.L.), and Cherry (#15 Sebewa loam and #16 Sloan loam). For all positive sites, three out of four had grass cover; i.e. two for loam (Min) and one for peat (Org). Only one site had leaf cover; i.e. loamy sand (Min). None had a mixture of grass and leaves. For all negative sites, one had grass cover; i.e. muck (Org). Two had leaf cover; i.e. sandy loam (Min) and five had a mixture of grass and leaves; i.e. sandy loam, loam (Min) and muck (Org). PERCENTAGE MOISTURE: The percentage moisture in different layers of all organic and mineral soils determined during the months of Octo- ber and November 1977 showed that the greatest amount of moisture was present at the litter layers of all sites, while lesser amounts were found at the 1" and 8" layers (Table 8). The highest percentage moisture recorded was 177% in the litter layers of #9 Fox loam (October) and #15 Sebewa loam (November). The lowest was 8% in the 8" layer of #15 Sebewa loam (November). The difference between percentage moisture from October to November ranged from 17 to 71% in the litter layers, 4 to 25% in the 1" layers and 2 to 9% in the 8" layers. This indicates that moisture in a given soil sample fluctuates widely within a short period of time (Table 9). All N. gruberi cysts and vegetatives isolates were obtained from sites with moisture levels between 17 and 35%. l8 BULK.DENSITY AND POROSITY: The bulk density measurements for the organic and mineral soils in some layers were found to be slightly higher than values obtained for these types of soils in other studies. This was due to many pieces of stones and rocks which had to be left intact within the samples. The percentage pore space for organic soils ranged from 68% in muck to 77% in peat, while mineral soils ranged from 39% in loamy sand to 50% in loam (Table 10). No major differences from normal range of porosity were observed. pH DETERMINATION: The pH of selected samples varied from 5.6 to 6.3 for organic soils (peat and muck) and 6.2 to 6.5 for mineral soils (loamy sand and loam). Therefore, both the organic and mineral soils tested were slightly acidic at all layers (Table 11). NAEGLERIA GROWTH IN THE SOIL: .Ngmbggs Per Gram Dry Soil: The numbers of Naegleria and Naegleria- like cysts and vegetative cells per gm of dry soil in different layers of positive sites during the months of August and September 1977 are shown in Table 12. Generally, more organisms were present in August than in September. There were more vegetative cells than cysts at most layers and more cells in soil layers than in litter layers. In litter layers, the highest number of cells per gm recorded were 6,111 (August, #7 Ottokee loamy sand) of which 1,524 were cysts; the lowest number was 159 (September, #10 Miami Loam) of which 98 were cysts. At the 1" layers, the highest number was 6,021 (August, #10 Miami loam) with 2,532 cysts; the lowest was 225 (September, #9 Fox loam) with 205 cysts. Finally, at the 8" layers, the highest number recorded was 16,288 (August, #9 Fox loam) with 2,618 cysts, while the 19 lowest number was 714 (September, #9 Fox loam) containing 326 cysts. Thus the numbers of organisms per gm in different layers varied from soil to soil, from layer to layer and from month to month. Both significant and non-significant differences were observed in both months at different layers (AppendixA and B). Growth in Autoclaved Soils: Inoculation of the #8 Peat (Org) and the #7 Loamy sand (Min) Naegleria isolates into 1 g of autoclaved soils from different layers of organic and mineral soils respectively in excess of population range in nature resulted in death of cells and lack of growth (Table 13). About 51,000 to 145,000 amoebae were inocu- lated per gm of soil. The percentage survival ranged from 1.04% in #15 Miami loam (Min) at the 8" layer to 29.72% in #8 Rifle peat (Org) at the 1" layer. The organisms did not exhibit preferential growth in positive sites over negative sites in autoclaved samples. Inoculation of these Naegleria isolates within population range in nature resulted in growth and increase in number of cells, although the increases were not significant (Table 14). About 4,000 amoebae per gm were inoculated. Growth percentages ranged from 7% in #8 Rifle peat (Org) at 8" layer to 80.25% in #15 Miami loam (Min) also at the 8" layer. Positive sites #8 Rifle peat (Org) and #7 Ottokee loamy sand (Min) were again not preferred over negative sites #6 Carlisle muck (Org) and #15 Miami loam (Min) in autoclaved samples. Growth in Unautoclaved Soils: Inoculation of #8 Peat (Org) and #7 Loamy sand (Min) Naegleria isolates into 1 g of unautoclaved soils from different layers of organic and mineral soils respectively with no bacterial food added resulted in significant growth in all layers 20 from positive sites and poor growth or death of cells in all layers from negative sites (Table 15). The highest growth percentages which were 187.68% and 245.18% occurred at the 1" layers of positive sites #8 Rifle peat (Org) and #7 Ottokee loamy sand (Min) respectively; the lowest growth percen- tages were -64.83% and -54.43% at the 8" layers of negative sites #6 Carlisle muck (Org) and #15 Miami loam (min) respectively. For all positive sites, the total number of cells after growth in unautoclaved soils far exceeded the total number in autoclaved soils by a wide margin. 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