Sulfur oxidation and respiration in 54-year-old soil samples

Sod Bml. Biochem. Vol. 9, pp. 405 to 410. Pergamon Press 1977. Prmted in Great Britam

SULFUR

OXIDATION 54-YEAR-OLD

AND RESPIRATION SOIL SAMPLES

WALTER USDA

B.

IN

BOLLEN

Forest Service, Pacific Northwest Forest and Range Experiment Forestry Sciences Laboratory, Corvallis, Oregon 97331, U.S.A.

Station.

(Accepted 12 March 1977) Summary-Soil samples in dry storage for 54yr were shown to retain their ability to respire and to oxidize S. Three of the soils had lower S-oxidizing capacity and three oxidized more S at 1 g kg-’ than did the samples when originally collected. When the experiment was repeated with all apparatus sterilized by autoclaving and S sterilized in flowing steam, a greater proportion of the S was oxidized. This was not due to heat treatment of the S. In all cases, S additions and incubation resulted in a lowering of the soil pH, suggesting that Thiohacillus thiooxidans was responsible and had survived the prolonged storage. When the soils, before and after incubation, were added to Thiobacillus media, only Gram-positive bacteria, mostly Bacillus spp., were found.

INTRODUCTION

Spore-forming and other bacteria, remain viable for long periods in dry soil and other harsh environments. Little is known concerning their potential ability to retain such metabolic activities as nitrification, sulfur oxidation, and organic matter decomposition during dormancy. Dry soil samples more than 15-year-old are capable of nitrifying added ammonium sulfate (Fraps and Sterges, 1932). Simms and Collins (1960) found that nitrifiers had strong resistance against drought in desert soils. In a 5-year study of dry zone soils in the Argentine Patagonia, Garbosky and Giambiagi (1962) observed that where Ca and K were adequate, initially high nitrification was maintained or only slightly decreased. In a study of S-oxidizing microorganisms in some Australian soils, including 23 desert sands and 21 desert loams, Vitolins and Swaby (1969) found that S was oxidized not only by autotrophic Thiobacilli but also by such heterotrophs as Arthrobacter aurescens, Bacillus licheniformis, and species of Fiavobacterium, Alcaligenes and Mycobacterium. Abbot (1923) showed that Penicillium luteum, another heterotroph, oxidized S to SO:-. Desert soils from Antarctica, Chile and California have been shown to have ammonifying, nitrifying and S-oxidizing potential (W. B. Bollen, K. M. Kemper, K. M. Byers and F. Au, unpublished reports). Although their specific metabolic activities were not determined, various bacteria have been cultured from desert soils of arid and frigid climates and from dried soil collections. Representatives of Bacillus, Mycobacterium, Mycoccus, Nocardia and Streptomyces were isolated by Bollen and Kemper (unpublished report) from samples of the Chile-Atacama desert, an area of very low and infrequent rainfall. Bollen and Byers (unpublished report) isolated species of Bacillus, soil coryneforms and Streptomyces from soil samples from the 70-year-old collection of E. W. Hilgard, late Professor of Soils at the University of California, Berkeley. Vela (1974) detected viable Azotobacter in soils stored in the laboratory 405

for more than 10 yr. I have observed that Azotobacter chroococcum can retain viability for 8-10yr in shrunken dried agar slope cultures that have become so hard that removal from the tube carries adherent flakes of glass. Species of Arthrobacter, related soil coryneform bacteria, Bacillus, Breuibacterium, Corynebacterium, Pseudomonas and Streptomyces constitute the major groups found in arid lands (Cameron, 1969). Such studies emphasize the remarkable ability of many bacteria to survive highly unfavorable environments for long times and to resume active life processes upon re-establishment of suitable conditions. In my study, certain stored samples of Oregon soils collected in 1921 for study of S oxidation (Halversen and Bollen, 1923) were tested again in 1975 and 1976 for retention of this property and also for soil respiration activity.

MATERIALS

AND

METHODS

The original 14 samples were collected in 1921 and represented four distinct areas of Oregon, but only six samples were available for this study: Columbia basin soils-Umatilla medium sand and Stanfield fine sand on which alfalfa responded to application of flour sulfur; southern Oregon soils-Salem clay loam, Salem clay loam which had been sulfured and Medford silt loam; Willamette Valley soil-Dayton silt loam. These air-dry samples had been stored in 946 ml fruit jars with clamped but unsealed lids, in the attic of a five-story building, subject to temperatures as low as - 12°C in winter and as high as 40°C in summer. Descriptions of the various samples are given by Halversen and Bollen (1923). Their chemical characteristics are presented in Table 1. Moisture was determined by drying at 105°C. Water-holding capacity was obtained from the weight of water retained by subsamples after saturation in large Gooch crucibles wetted from below and then drained to constant weight in a saturated atmosphere. For pH, a glass electrode was used on 1: 5 w/v

at 112 kg ha at l12kgha-’

characterization

in 1917.

31.1 40.1

28.1 29.4 38.7 40.7

Water-holding capacity (“0)

I. Chemical

1919. acre in 1915, and again

I in March

1.80 2.05

Medford silt loam Dayton silt loam

* Oven-dry basis. t Flour sulfur applied 8 Flour sulfur applied

0.97 1.14 3.60 3.64

Umatilla medium sand Stanfield fine sand, sulfuredt Salem clay loam Salem clay loam, sulfured:

(““I

Water

Table

6.7 5.6

6.8 6.8 6.6 6.8

PH

of w-dry

93.29 94.30

97.86 97.16 95.38 93.48

Ash (“@

54-year-old

1.62

2.03

0.47 0.67 I .86 1.95

&

Total

soil samples*

0.188 0.118

0.047 0.083 0.132 0.151

G

Kjeldahl

10.80 13.61

10.00 8.07 14.09 12.91

C:N ratio

80 164

55 110 155 135

Total S parts/ 1Oh

6 I

s 38 8 8

so:-ms parts /IOh

co ; r L

g Y L 7 f F

407

Sulfur oxidation and respiration in 54-year-old soil samples aqueous soil suspensions while stirring. Soil analyses were made on subsamples ground to pass a 149 pmmesh sieve. Burning in a muffle furnace at 650°C provided data on ash and loss on ignition. Total N was determined by Kjeldahl procedure and total C by dry combustion at 950°C (Allison et al., 1965). Total S was determined in 1921 by fusion of 2g soil with 14 g sodium peroxide, 0.8 g magnesium powder and 1 g sucrose in an illium sulfur bomb (Parr Instrument Co., Moline, Illinois). Subsequent procedure was as recommended by the Association of Official Agricultural Chemists (1920). Sulfates were determined turbidimetrically (American Public Health Association, 1955) on water extracts clarified by passage through Millipore 0.22 pm filters. Analytical data for these soils are in Tables 1 and 2. Numbers of molds were determined by plating dilutions with peptone glucose acid agar; bacteria and streptomyces, with sodium albuminate agar (Fred and Waksman, 1922); and endospores, by plating dilutions of pasteurized 1:5 (w/v) soil suspensions with nutrient agar. For thiobacilli (Vishniac and Santer, 1957), inorganic media adjusted to pH 3.5 for ThiobaciUus thiooxidans and to pH 6.8 for 7’. thioparus were used. To detect acid production, 0.25% tri-calcium phosphate was added. A clear zone around a colony indicated dissolving of the insoluble salt. All plates were incubated at 28°C. The study comprised two separate experiments. The first was conducted with the usual precautions and cleanliness employed in soil microbiology, but apparatus and containers were not sterilized. Each soil and each treatment was run in duplicate and in random order, but the replications were run at different times over a period of 16 months. For S oxidation

and soil respiration, 1OOmg flour sulfur was mixed with 1OOg soil while 100 g untreated soil was used as a control. The soils were placed in 473-ml bottles and distilled water added to adjust moisture content to 50% of water-holding capacity. The bottles were then connected to a manifold supplying air washed through l~Na0H and saturated Ba(OH), solution to remove COZ, through 1~ H2S04 and through water to maintain soil moisture. Each bottle outlet was connected to a sparger immersed in a test tube containing 10ml 1~ NaOH. Carbon evolved as CO2 was absorbed by the NaOH. At 7, 14, 21 and 28 days after treatment, the tubes of alkali were replaced and CO2 determined by differential titration with 1~ and 0.08~ HISO, using a Beckman automatic titrator and Cooper’s (1941) end points. Analyses for sulfates and pH values were made on the soil in each bottle at the end of the 28-day incubation. To avoid any contamination with S-oxidizing organisms, the second experiment was conducted with aseptic sampling and all equipment sterilized by autoclaving or flowing steam. Because Waksman (1922) believed that 7: thiooxidans are usually not originally present in the soil but are introduced artificially with the S added, the S (precipitated sulfur) used in this experiment was placed in a 250-ml Erlenmeyer cotton plugged flask and sterilized in flowing steam for 30min on three consecutive days. This S gave no indication of any growth within 30 days at 28°C when tested on Waksman’s (1922) liquid and solid media and the Vishniac and Santer (1957) medium. It was found that the unsterilized precipitated S, as well as three different brands of agricultural flour sulfur tested in a similar manner, did not carry thiobacilli. The setup was similar to the first

Table 2. Microbial analyses of soils (numbers per gram, oven-dry basis) Molds Soil No.

Thousands

Total lo6 g- ’

S/lo6

> lO$ >500 16.4

4

Original sample Control 1000 parts S/lo6

> 10 3.7 1.5

0.2 31.3 39.2

5

Original sample Control 1000 parts S/lo6

> 10 17.0 1.0

6

Original sample Control 1000 parts S/lo6

7

13

103g-’

Treatment

Endospores

Thiobacteria* at pH 6.8

at pH 3.5

(%I

103g-’

0.2

5.0

110

11.4 21.6

6.5 3.5

20 4925 1400

100 1163 2660

3.0 6.0 3.5

120

70 6045 2125

90 5225 5280

0.4 30.1 48.0

13.1 23.0 5.2

710

370 3515 6265

1120 4275 4315

0.2 4.1 5.8

1.0 32.4 45.5

7.9 16.5 8.3

400

120 4100 6245

700 6700 5325

Original sample Control 1000 parts S/lo6

> 10 16.3 11.3

1.1 38.6 43.0

3.5 14.5 10.9

420

90 4800 3005

1000 3225 718

Original sample Control 1000 parts S/lo6

> 10 22.9 20.7

0.2 9.3 2.6

1.8 9.0 1.5

420

710 4775 4135

710 333 205

Original sample7 1

Bacteria Streptomyces

Control$ 1000 parts

* Isolated on Vishniac and Santer (1957) medium. Not Thiobacillus spp. t Air dry. Not incubated. $ > indicates no growth on 1: 10 or 1: 500 dilutions. 6 Incubated 28 days at 28°C with moisture at 50% of water-holding capacity sterile apparatus and aseptic procedure.

and

with

continuous

aeration;

all

WALTER B. BOLLFR

10x

run except that the brass manifold was steam sterilized and the CO,-free air was passed through additional 1~ H,SO, and sterile distilled water, then over VishniaccSanter (1957) medium and nutrient agar before entering the manifold. These media remained sterile during the course of the experiment, To check for any H,S or S that could possibly be derived from rubber tubing and stoppers used in the connections, suitable traps were attached to the manifold; no S, sulfide, or sulfate was detected. In Table 3, the second experiment is referred to as “aseptic” while the first is designated “clean”. At the end of 28 days incubation in the second experiment, dilutions of the soils were plated for molds, bacteria and S oxidizers using the media previously mentioned. Plate counts were not made in the first experiment. All data presented are means of duplicate determinations. RESULTS

AND DISCUSSION

Few molds and bacteria were found in the original air-dry samples and a large proportion of the bacteria were spore formers (Table 2). Each of the old soil samples retained capacity to oxidize S with marked lowering of pH (Table 3). Under the usual testing conditions (apparatus not sterilized), three of the soils oxidized less of the I g kg-’ added S than in the 1922 study; three oxidized more. With sterilized apparatus, however, all the soils with added S produced more sulfate. Only in Dayton silt loam was the difference between runs with clean and sterilized apparatus minor. In two cases, sulfate recovery slightly exceeded IOO”~,,indicating either an analytical error due to the very high dilution required. or perhaps additional sulfate from oxidation of some soil organic S. Sulfate found in the controls did not greatly differ between the clean and sterile apparatus usage. Why more S was converted to sulfate with the sterile apparatus is not clear. Possibly more aeration induced by mixing and additional exposure in taking samples for the second experiment could be a factor. Repeating the aseptic study with the S not sterilized by flowing steam gave similar results, showing that the heat treatment did not render the S more susceptible to oxidation. Whether or not T. tkioosidnns or other species of bacteria were responsible for the oxidation was not satisfactorily determined. Microbial analysis were made on the original soil samples and soils incubated in aseptic apparatus. In addition to media for hetertrophs. samples were also plated in I : 10 and I :50 dilutions with Thiohcrcillus medium (Vishniac and Santer. 1957) modified by 0.2”,, tricalcium phosphate in suspension. One series of plates was poured with the medium adjusted to pH 3.5 for T. thioo.uidans and another adjusted to pH 6.8 for T. (hioporus and lower acid-producing species. All the samples except No. 13 gave generally higher counts on the less acid medium. Incubation of the original samples increased their colony counts (Table 3); with added S, the increases in total bacteria were much greater except for Dayton silt loam; numbers of Struptomycrs, on the other hand, were reduced in all cases. Colonies on the Thiohaci2lu.s media were small, about 2-mm dia, dark cream turning brown in old

cultures, dull and opaque. On the medium adjusted to pH 3.5, clear zones, l- to 2-mm wide, indicating solution of the tricalcium phosphate, were developed with soils 5, 6 and 7 but not with the others. Bacteria in these soils produced higher acid concentrations. In no case did the organisms prove to be T/n&acillus. Gram stains of smears from colonies revealed only Gram-positive rods, 1.221.5 x 335 pm, single and in chains. sometimes in long filaments. Endospores appeared in non-swollen sporangia. Transfers to nutrient agar grew well. It seems that Bacillus sp. may be the S-oxidizing agent in the old soil samples. Studies are in progress to characterize the organism and determine its capability for oxidizing sulfur in pure culture. R. J. Swaby (personal communication) finds no correlation between the incidence of T. fhiooxi&ra.s and the capacity of a soil to oxidize S; but when this species is abundant. then oxidation is usually rapid. It can still be rapid due to other T/r&hrrcillus species. Certain heterotrophs can oxidize S (Vitolins and Swaby, 1969). but R. J. Swaby (personal communication) believes they are relatively unimportant in soils rapidly carrying on the oxidation. I. L. Pepper (personal communication) recently isolated from soil a heterotrophic Micrococc~rs capable of oxidizing elemental S to SO: The very rapid oxidation of S in the Stanfield soil suggests that T. thioo.xir/nns was responsible. and the application of S to the field in 1919 could have built up the population before the sample was collected in 1921. This does not hold for the Salem soil, where the previous sulfuring gave no advantage over the unsulfured soil. In both cases. the S oxidized in the 54-year-old samples was a moderate S’,,. Vitolins and Swaby (1969) using l”,, S additions and incubation for 10 weeks at 25 C. considered the amount of oxidation to be high at 13 to 60”,, and moderate at 4 to 13”,,. Based on these criteria and adjusting them to 1 g S kgg ’ additions and incubation for 28 days, two of the soils in Table 2 show a high rate of S oxidation while the others are in the moderate range. Other species could be responsible for retention of the capacity for S oxidation. but their persistence for 54yr in the dry soils seems remarkable unless some spore-forming species were the active agents. Vitolins and Swaby (1969) believe the role of any heterotrophs, either cyst- or spore-formers, is remote with moderate to high S oxidizing rate and low pH values. Our failure, however, to isolate Thiohucillus from any of the soils, and finding highly acid-tolerant Baci//w in all. points to some member of this spore-forming genus as the S-oxidizer. Organic matter decomposition, as indicated by CO1 production (Table 3). was within the ranges commonly found for may soils. The extensive decomposition shows the Stanfield soil is unusual, considering the low C content. Despite low organic matter contents. many sandy and pumice soils have been observed to evolve CO2 more rapidly than finer textured soils much higher in organic matter. This could be attributable to better aeration. Although less remarkable than the persistence of S oxidizing capacity, the ability of the old, dry soils to actively evolve CO1 when remoistened and incubated is noteworthy. Colonies of Bucillus species predominated in samples of the dry soils sprinkled di-

21 23

22 29

49 20

29 26

5.6 5.7

5.8 5.9

7.0 6.9

5.9 5.9

clean aseptic

clean aseptic

clean aseptic

clean aseptic

5

6

7

13

4.9 5.3

6.3 5.2

5.1 4.9

4.6 4.4

4.6 4.1

4.2 4.2

84 97

113 123

times during

124 348

55 1021

52 440

789 1062

52 500

parts/l06

163 368

17 1050

73 463

860 1166

85 513

parts/l06

samples

incubated

16 months.

8.4 9.7

12.4 34.8

5.5 100.0

5.2 44.0

78.9 100.0

5.2 50.0

%

46.7 66.2

85.4 69.2

11

2

63.3 63.1

63.7 61.6

7

10

52.2 29.4

21.3 40.6

mg 100 g-r

40.7 47.9

2.88 4.09

4.21 3.41

3.25 3.24

51.8 59.3 81.8 66.4

3.42 3.31

7.79 4.39

4.53 8.63

%

2.51 2.96

4.03 3.27

2.66 3.04

2.84 2.74

7.52 3.19

3.85 4.60

%

Soil C oxidized to CO2 1000 parts S/lo6 control

53.9 51.0

50.4 21.4

S/lo6

procedures

18.1 21.6

soil

as CO2 1000 parts

clean vs. aseptic

C evolved control

28 days*?,

38

28

%

S oxidized in 1975-1976 in 1922 28 days in 14 days1

and CO2 efflux from 55year-old

* At 28°C with moisture adjusted to 50% of water-holding capacity. t Data are means of duplicate runs made in random order at different $ Halverson and Bollen, 1923.

81 104

7.3 6.4

clean aseptic

4

33 13

6.7 6.5

parts/lo6

clean aseptic

Apparatus used

1

Soil No.

3. Sulfur oxidation

pH and S as SOicontrol 1000 parts S/lo6 pH SO:--S pH so:-PS

Table

WALER

410 rcctly

onto

nutrient

agar plates. These bacteria

could

be largely responsible With

each

for the CO2 production. soil, the S addition in the clean experi-

ment decreased CO, evolutionPprobably due to the increase in acidity (Table 3). Organic matter decomposition typically proceeds less rapidly as pH decreases. On the basis of soil total C, the decrease was greatest in the Umatilla soil, which had the lowest C content, and least in the Medford soil, which had the most C. In the second experiment, with sterile apparatus and sterile air, CO2 evolution, with two exceptions, was greater than in the clean setup. In only two cases was more CO2 evolved from controls than from the S-treated soils. Revival of these activities by bacteria that have survived and remained potent for prolonged periods in dry soils raises questions concerning contributing factors. Answers will require investigation of the physico-chemical and biological conditions and processes involved. The energy relations of water retention and exchange between an air-dry matrix and cells existing on hygroscopic films require investigation.

REFERENCES

ABBOTT E. V. (1923) The occurrence and action of fungi in soils. Soil Sci. 16, 207--216. ALLISON L. E., BOLLEN W. B. and MOODIE C. D. (1965) Total carbon. Methods of’ Soil A&@~, Part 2, pp, 1356-1360. Am. Sot. Apron. Madison, Wisconsin. AMERICAN PUBLIC HEALTH ASSOCIATION (1955) Standard

B. BOLLEN Methods for the Examination of Water. Sewage and Industrial Wastes, pp. 197-198. New York. ASSIXIATION OF OFFICIAL AC;RICULTURAI. CHEMISTS (1920) Oficicrl Merkods. Washington. D.C. CAMERON R. E. (1969) Cold desert characteristics and problems relevant to other arid lands. In 4rid Lads in PYYspecricr (William G. McGinnies and Bram J. Goldman, Eds), pp. 169-205. Atn. .4ss. .4i/r. Ser. Univ. Arizona Press, Tucson. COOPER S. C. (1941) The mixed indicator bromscresol green-methyl red for carbonates in water. I&. Engng. Chem. anal@ Edn. 13, 466470. FRAPS G. S. and STERGPS A. J. (1932) Causes of low nitrlfication capacity of soils. Soil Sci. 34, 353-363. FRED E. B. and WAKSMAX S. A. (1922) A tentative outline of the plate method for determining the number of microorganisms in the soil. Soi( Sci. 14, 27-28. GARBOSKY A. J. and GIAMBIAC;I N. (1962) The survival of nitrifying bacteria in the soil. PI. Soil 17, 271-278. HALVERSEN W. V. and BOLLEN W. B. (1923) Studies on sulfur oxidation in Oregon Soils. Soil Sci. 16, 479-490. SIMS C. M. and COLLINS F. M. (1960) The numbers and distribution of ammonia oxidizing bacteria in some northern territory and South Australian soils. 4ust. .I. agric. Res. 11, 505-512. VELA G. R. (1974) Survival of Azotohtrc,fer in dry soil. Appl. Microhiol. 28, 77 79. VISHNIAC W. and SANTER M. (1957) The thiobacilli. Bact. Rev. 21, 195-213. VITOLINS M. I. and SWABY R. J. (1969) Activity of sulphuroxidizing microorganisms in some Australian soils. Aust. J. Soil Rex 7, 171-183. WAKSMAN S. A. (1922) Microorgamsms concerned in the oxidation of sulfur in the soil, IV. A solid medium for the isolation and cultivation of Thiohacillus thiooxidnns. J. Burt. 7, 605%608.