letters to nature
Acknowledgements
The calculations were performed using the NERC-supported Cray T3E machines at the
Manchester CSAR Centre and at the Edinburgh Parallel Computer Centre, and the UCL
HiPerSPACE Centre. Support from the NERC and discussions with L. VocÆadlo are
acknowledged.
Correspondence and requests for materials should be addressed to D.A.
(e-mail: d.alfe@ucl.ac.uk).
.................................................................
Detection and classi®cation
of atmospheric methane
oxidizing bacteria in soil
Ian D. Bull*, Nisha R. Parekh², Grahame H. Hall³, Philip Ineson§
& Richard P. Evershed*
* School of Chemistry, University of Bristol, Cantock's Close, Bristol BS8 1TS, UK
² Institute of Terrestrial Ecology, Merlewood Research Station,
Grange-over-Sands, Cumbria LA11 6JU, UK
³ Institute of Freshwater Ecology, Far Sawrey, Ambleside, Cumbria LA22 0LP, UK
§ Department of Biology, University of York, P.O. Box 373, York YO10 5YW, UK
..............................................................................................................................................
Well-drained non-agricultural soils mediate the oxidation of
methane directly from the atmosphere, contributing 5 to 10%
towards the global methane sink1,2. Studies of methane oxidation
kinetics in soil infer the activity of two methanotrophic populations: one that is only active at high methane concentrations (low
af®nity) and another that tolerates atmospheric levels of methane
(high af®nity). The activity of the latter has not been demonstrated by cultured laboratory strains of methanotrophs, leaving
the microbiology of methane oxidation at atmospheric concentrations unclear3,4. Here we describe a new pulse-chase experiment using long-term enrichment with 12CH4 followed by shortterm exposure to 13CH4 to isotopically label methanotrophs in a
soil from a temperate forest. Analysis of labelled phospholipid
fatty acids (PLFAs) provided unambiguous evidence of methane
assimilation at true atmospheric concentrations (1.8±3.6
p.p.m.v.). High proportions of 13C-labelled C18 fatty acids and
NATURE | VOL 405 | 11 MAY 2000 | www.nature.com
the co-occurrence of a labelled, branched C17 fatty acid indicated
that a new methanotroph, similar at the PLFA level to known type
II methanotrophs, was the predominant soil micro-organism
responsible for atmospheric methane oxidation.
Culturable methane oxidizing bacteria are classi®ed into types I,
II and X depending on the guanine and cytosine content of their
DNA, intracellular membrane arrangement, carbon assimilation
pathway and PLFA composition3. Active soil microbial populations
utilizing a 13C-labelled substrate will readily incorporate 13C into
membrane lipid components such as PLFAs. Recent work has
demonstrated the power of 13C-labelling to link lipid biomarkers
to other biogeochemical processes in lake and marine sediments5,6.
We have applied similar techniques to a soil to investigate the
oxidation of methane at atmospheric concentrations.
Soils used for pulse-chase labelling were obtained from a Sitka
Spruce plantation in North Wales, UK. Mull-board ploughing
before tree planting in 1959 produced a repeating microtopographic
pattern consisting of a plough furrow and ridge followed by an
undisturbed ridge. The plough ridges have an inversion of the
original strati®cation of soil horizons resulting in a buried organic
horizon beneath an overturned mineral horizon; soil cores from
these ridges exhibited high rates of atmospheric methane consumption (mean rate is 77 mg m-2 h-1, standard error (s.e.) = 2, n = 7).
Laboratory experiments at atmospheric and elevated concentrations demonstrated that soils from both the buried organic and
mineral horizons showed similar kinetics of oxidation, but activities
were generally lower in the mineral soil. Increasing the initial
concentration of methane from 1.8 to 300 parts per million by
volume (p.p.m.v.) for both soils showed typical Michaelis±Menton
kinetics with Km methane concentration at half of the maximal
reaction rate values of 25 and 56 p.p.m.v. for the buried organic and
mineral soils, respectively (Fig. 1). Rapid rates of methane consumption, characteristic of high-af®nity methane-oxidation
kinetics, were observed in both soils below 50 p.p.m.v. methane.
Concentrations of methane, which induce oxidation kinetics indicative of high (1.8±3.6 p.p.m.v.) and low (100 p.p.m.v.) af®nity
methanotroph populations, were chosen for the 13C pulse-chase
experiment in soil columns.
The distribution of PLFAs shown in Fig. 2 (for the buried organic
horizon) is typical of all the soil samples analysed; the only
signi®cant difference between the two soil types is the lower overall
abundance of PLFAs in the mineral soil (,35 mg g-1
TOC); where TOC
refers to `total organic carbon' compared with the buried organic
Rate of oxidation (nmol CH4 per gram
dry weight of soil per hour)
15. de Wijs, G. A., Kresse, G. & Gillan, M. J. First-order phase transitions by ®rst-principles free-energy
calculations: The melting of A1. Phys. Rev. B 57, 8223±8334 (1998).
16. AlfeÁ, D., Gillan, M. J. & Price, G. D. The melting curve of iron at the pressures of the Earth's core from
ab initio calculations. Nature 401, 462±464 (1999).
17. AlfeÁ, D., Gillan, M. J. & Price, G. D. Thermodynamics of hexagonal-close-packed iron under Earth's
core conditions. Phys. Rev. B (submitted).
18. AlfeÁ, D., de Wijs, G. A., Kresse, G. & Gillan, M. J. Recent developments in ab-initio thermodynamics.
Int. J. Quant. Chem. 77; 871±879 (2000).
19. Giannozzi, P., de Gironcoli, S., Pavone, P. & Baroni, S. Ab initio calculations of phonon dispersions in
semiconductors. Phys. Rev. B 43, 7231±7242 (1991).
20. Kresse, G., FurthmuÈller, J. & Hafner, J. Ab initio force-constant approach to phonon dispersion
relations of diamond and graphite. Europhys. Lett. 32, 729±734 (1995).
21. Chandler, D. Introduction to Modern Statistical Mechanics (Oxford University Press, 1987).
22. Frenkel, D. & Smit, B. Understanding Molecular Simulation Ch. 4 (Academic, New York, 1987).
23. Sugino, O. & Car, R. Ab initio molecular dynamics study of ®rst-order phase transitions: melting of
silicon. Phys. Rev. Lett. 74, 1823±1826 (1995).
24. Kresse, G. & FurthmuÈller, J. Ef®cient iterative schemes for ab initio total-energy calculations using a
plane-wave basis set. Phys. Rev. B 54, 11169±11186 (1996).
25. VocÆadlo, L., Brodholt, J., AlfeÁ, D., Price, G. D. & Gillan, M. J. The structure of iron under the
conditions of the Earth's inner core. Geophys. Res. Lett. 26, 1231±1234 (1999).
26. Usselman, T. M. Experimental approach to the state of the core: part I. The liquidus relations of the Ferich portion of the Fe-Ni-S system from 30 to 100 kb. Am. J. Sci. 275, 278±290 (1975).
27. Hammond, B. L., Lester, W. A. Jr & Reynolds, P. J. Monte Carlo Methods in Ab Initio Quantum
Chemistry (World Scienti®c, Singapore, 1994).
28. Rajagopal, G., Needs, R. J., James, A., Kenny, S. D. & Foulkes, W. M. C. Variational and diffusion
quantum Monte Carlo calculations at nonzero wave vectors: Theory and applications to diamondstructure germanium. Phys. Rev. B 51, 10591±10600 (1995).
29. Kent, P. R. C. et al. Finite-size errors in quantum many-body simulations of extended systems. Phys.
Rev. B 59, 1917±1929 (1999).
30. Kilburn, M. R. & Wood, B. J. Metal-silicate partitioning and the incompatibility of S and Si during
core formation. Earth Planet. Sci. Lett. 152, 139±148 (1997).
8
6
4
2
0
0
50
100
150
200
Initial [CH4] (p.p.m.v.)
250
Figure 1 Rates of methane oxidation between 1.5 and 250 p.p.m.v. by forest soil from the
mineral and buried organic layers. Lines represent direct nonlinear regression analysis
and show typical Michaelis±Menton kinetics. Data points show the mineral layer (white
circles) and buried organic layer (black circles). Application of the Lineweaver Burke plot
provided a signi®cant regression (P , 0.001) for both data sets that were used to
calculate the apparent Km (methane concentration at half of the maximal reaction rate) and
Vmax (maximal reaction rate) values for each soil. Regression for mineral soil R2 = 0.78,
apparent Km = 56 p.p.m.v., apparent Vmax = 2.8 nmol per gram dry weight of soil per hour.
Regression for organic soil R2 = 0.82, apparent Km = 25 p.p.m.v., apparent Vmax = 3.7
nmol per gram dry weight of soil per hour.
© 2000 Macmillan Magazines Ltd
175
letters to nature
strains, although not the species Methylosinus trichosporium, could
possibly account for the heavily labelled 18:1v7c component, the
lack of labelled 18:1v8c and the low number of other 13C labelled
PLFAs8,9. Indeed, this would be entirely consistent with previous
work which has utilized gene cloning and sequencing techniques
to identify the presence of this phylogenetic group in a four-yearlong liquid enrichment culture under elevated concentrations of
methane (,275 p.p.m.v.; ref. 10). However, the co-occurrence of an
equally labelled br17:0 PLFA in the soil samples is not consistent
with the PLFA pro®les of previously described type II methanotrophs. Trace amounts of a br17:0 component have been detected in
Methylobacterium organophilum and Methylosinus trichosporium as
well as in natural-gas-enriched soils7±9,11,12. The data shows that
although the br17:0 component constitutes a small proportion of
the total PLFAs in all of the methane-treated soils (6.4±7.8%), the
degree of incorporation of carbon from 13CH4 in this component is
extremely high compared with that in the 16:1 and 18:1 signature
lipids for known methanotrophic bacteria. This supports the
proposition that methane oxidation is mediated predominantly
by a new methanotroph which expresses some similarity at the
PLFA level to known type II methanotrophs.
The buried organic soil exposed to elevated levels of methane
(100 p.p.m.v.) exhibits signi®cant 13C-labelling of the 16:0, 16:1v8c,
a
350
300
250
200
δ13C (‰)
soil (,75 mg g-1
TOC). This quantitative difference has been reported
previously and arises from the inherently higher biomass sustained
by the nutrient-rich buried organic soil7. This is also re¯ected in the
higher rates of methane oxidation, and therefore presumably greater
biomass, in the buried organic horizon. PLFAs in both the buried
organic and mineral soils comprise mainly C16 and C18 fatty acids
and apart from the saturated components are composed predominantly of 1v7 (indicative of Gram negative aerobes, obligate anaerobes) and 1v9 (indicative of Gram positive bacteria) unsaturated
compounds8. Signi®cant quantities of 18:1v8c unsaturates, considered highly indicative of type II methanotrophs such as
Methylosinus trichosporium, are not detectable in any of the soils
(Fig. 2). Although used routinely for studies of soil biomass and
bacterial community structure, the drawbacks of quantitative lipid
pro®ling are quickly realized when attempting to assess the microbial populations involved in speci®c processes within the same soil.
No signi®cant differences can be detected between the PLFA
compositions of soils exposed to 1.8±3.6 p.p.m.v. methane and
those incubated under elevated methane (100 p.p.m.v.; results not
shown) because methanotrophs are minor contributors to the
overall soil microbial biomass. However, incorporation and subsequent detection of a stable isotope label by a pulse-chasing
approach using a process substrate, for example methane (13CH4),
easily overcomes limitations associated with the pro®ling method.
In this way gas chromatography combustion isotope ratio mass
spectrometry provides a uniquely speci®c and sensitive means of
determining any metabolic incorporation of the 13C label into
PLFAs derived from populations of methane-oxidizing bacteria.
Analysis of PLFAs from the buried organic soil exposed to
atmospheric levels of methane (1.8±3.6 p.p.m.v.) clearly shows
incorporation of the 13C label in the br17:0 (component 16) and
18:1v7c (component 27) PLFAs (Fig. 3a). This indicates that the
oxidation of atmospheric methane in this soil is mediated primarily
by organisms with a labelling pattern similar to that of a type II
methanotroph8,9. The presence of type II Methylosinus/Methylocystis
150
100
10
50
26
0
27
–50
1
Relative intensity
b
25
6
3
8
9
2
4
10
12
7
11
12
18
13
14 19
21
15
16
22
17
20 23
16
14
Retention time (min)
200
29
28
18
150
100
20
50
Figure 2 A partial gas chromatogram of the methylated phospholipid fatty acid (PLFA)
fraction derived from the buried organic soil exposed to an atmospheric (1.8±3.6
p.p.m.v.) concentration of methane. Peak numbers indicate PLFAs as follows: peak
number 1 is 12:0; 2 is nonadecane (internal standard; IS); 3 is 14:0; 4 is 8:0 diacid; 5 is
i15:0; 6 is a15:0; 7 is 15:0; 8 is 9:0 diacid; 9 is br16:0; 10 is 16:0; 11 is 16:1v11; 12 is
16:1v8c; 13 is 16:1v7c; 14 is br17:0; 15 is 16:1v5t; 16 is br17:0; 17 is i17:0; 18 is
a17:0; 19 is br17:1; 20 is br18:0; 21 is 17:0; 22 is 17:1v8; 23 is br18:0; 24 is 18:0; 25
is 18:1v13; 26 is 18:1v9c; 27 is 18:1v7c; 28 is 18:1v5c; 29 is 18:2; where br/a/i
indicates a branched/iso/anteiso alkyl chain; the number preceeding a colon indicates the
total carbon number; the number after the colon indicates the number of double bonds;
the number following v indicates the position of unsaturation counting from the terminal
methyl group; and c or t indicates a cis or trans geometric isomer.
176
PLFA
300
250
24
δ13C (‰)
5
3 5 6 10 11 12 13 14 15 16 17 19 20 21 23 24 26 27 28
0
–50
3 5 6 10 11 12 13 14 15 16 17 19 20 21 23 24 26 27 28
PLFA
Figure 3 A plot of d C values of PLFAs from soils after incubation and 13CH4 pulselabelling. a, Three triplicate buried organic soils. b, Three triplicate mineral soils. Pulselabelling was done under elevated methane (100 p.p.m.v., white squares), atmospheric
methane (1.8±3.6 p.p.m.v., black triangles) and control (non-enriched and unlabelled
soil, black circles). Missing data points correspond to peaks for which a reliable
measurement could not be obtained. Numbers on the x axes correspond to the PLFAs
listed in the caption to Fig. 2.
© 2000 Macmillan Magazines Ltd
13
NATURE | VOL 405 | 11 MAY 2000 | www.nature.com
letters to nature
16:1v7c, br17:0, br18:0, 18:1v9c and 18:1v7c components (Fig. 3a).
The range is limited and indicates an absence of any signi®cant
recycling and uptake of 13C through 13CO2 by autotrophic bacteria
during the course of the experiment. Such high levels of incorporation in both C16 and C18 PLFAs suggest that low-af®nity methane
oxidation in this soil is being performed by organisms related to
both type I and type II methanotrophs11. In addition, the small
number of heavily labelled components indicates that this process is
still, even under high concentrations of methane, being mediated by
a limited diversity of bacterial species. The labelled type I signature
(16:1) PLFAs exhibited by the soil are closely matched by those
present in Methylomonas gracilus, indicating that either this, or a
closely related species, may be partially responsible for low-af®nity
methanotrophy in this soil12. When soil is exposed to elevated
concentrations of methane, the activity of low-af®nity species will
increase and the activity of high-af®nity species will continue.
Therefore, the same type II species operating at atmospheric
concentrations of methane are likely to constitute a signi®cant
proportion of the methanotrophic population in the soils exposed
to elevated methane13.
Stable isotope analyses of individual PLFAs from mineral soil
exposed to both atmospheric (1.8±3.6 p.p.m.v.) and elevated (100
p.p.m.v.) levels of methane show incorporation of the 13C-label in
the 16:0, br17:0 and 18:1v7c PLFAs (Fig. 3b). Comparison with the
pro®les of reference bacteria suggests a population of methanotrophs dominated by an unknown organism, which is possibly
related to the type II Methylosinus/Methylocystis genera8. This
contrasts with the active methanotrophic population in the
buried organic soil exposed to elevated methane, which included
organisms related to both type I and type II methanotrophs. A
similar population of high-af®nity methane-oxidizing bacteria,
dominated by new organisms related to type II genera, are present
in both the buried organic and mineral horizons of this forest soil.
The ability to characterize active populations of bacteria in situ at
true atmospheric concentrations of methane facilitates a reliable
and ecologically relevant understanding of soil methane oxidation.
Alternative DNA sequencing techniques lack sensitivity and require
arti®cially high concentrations of methane10, whereas radiocarbon
labelling with 14C is also compromised by a lack of both sensitivity
and compound speci®city14. Compound-speci®c stable-isotope
analysis provides a sensitive and non-hazardous means of investigating the activity and roles of microorganisms involved in important biogeochemical cycles. In addition, biologically relevant
signature compounds, other than PLFAs, may be studied (for
example, non-standard amino acids, amino sugars, hopanoids,
ergosteroids and mycolipids). Information about soil methanotrophs can be used to formulate land management strategies with
the long-term goal of controlling and maximizing the quantity of
methane oxidized by terrestrial biomes.
M
Methods
Methane-oxidation potentials
Soil cores, of internal diameters 15 and 30 cm, were collected from plough ridges on the
¯oor of a temperate Sitka Spruce forest called Coed Anafon situated on the North Wales
Coast (538129 N, 48009 W). The soil cores were separated into three horizons; the upper
organic, the mineral , and the buried organic horizons. Samples of both the buried organic
and mineral soil horizons were sieved (2-mm mesh) and stored at 4 8C until required (,6
months). For the kinetic studies, the soil samples were acclimatized overnight at 20 8C,
thoroughly mixed and triplicate 10-g samples placed in serum bottles to which 10 ml of
sterile water was added. The bottles were sealed using butyl rubber septa and aluminium
crimps. Incubation was at 20 8C on a shaking table set at 150 r.p.m. The rate of methane
oxidation was estimated from the decrease in headspace concentration over 60 to 120 min
to ensure that instantaneous rates were determined. The short incubation measures the
activity of methane monooxygenase enzyme present in the sample and not the activity that
develops after exposure to added methane. At least three samples of headspace gas were
removed during the incubation period. The methane concentration in the headspace was
changed by adding standard methane/air gas mixtures. The concentration range for each
soil horizon was varied from 1.7 to 8,000 p.p.m.v. methane. Methane concentrations were
determined by gas chromatography15.
NATURE | VOL 405 | 11 MAY 2000 | www.nature.com
13
C-methane labelling
Glass sinter tubes with 20 g dry weight equivalent of soil from the buried organic or
mineral horizons were used as microcosms and incubated in a 20 8C constant temperature
room. A peristaltic pump was used to ¯ow methane in air, at atmospheric (1.8 p.p.m.v.) or
elevated (100 p.p.m.v.) concentrations, and sterile distilled water through the microcosms
at constant rates of 0.05 ml CH4 min-1 and 0.015 ml H2O min-1, respectively, over a sixmonth incubation period. The microcosms were sealed only at the top to allow for
drainage and venting of excess gas. The headspace gases over the equilibriated soils in the
atmospheric and elevated methane microcosms were then respectively replaced with 1.8 or
100 13CH4 in air. Microcosms were incubated for a further three weeks to label the active
methanotrophic bacteria populations within these soils. Thus, the microcosms used to
simulate atmospheric methane oxidation contained no more than 3.6 p.p.m.v. methane at
any stage of the experiment. 13CH4 was obtained from Isotech Inc., USA and graded 99.9%
pure and 99.4% 13C atom enriched. At the end of the equilibration and 13C-labelling
period, sub-samples of each soil were freeze-dried and PLFAs were extracted from these
samples.
PLFA analysis
Lipids were extracted by following a modi®ed Bligh and Dyer method16,17. Subsequent
fractionation on a silica column with a bonded amino-propyl phase yielded a total PLFA
fraction. Fatty acid components were released by mild alkaline hydrolysis (0.1 M KOH in
MeOH) and then isolated and methylated using a 14% (w/v) solution of a borontri¯uoride±methanol complex. Individual compounds were quanti®ed using an HP 5890
series II gas chromatograph equipped with a Chrompack CPWax-52CB (50 m length,
0.32 mm internal diameter, 0.12 mm ®lm thickness; H2 carrier gas, 10 psi head-pressure).
Compound structures were assigned unambiguously by gas chromatography/mass spectrometry through the analysis of dimethyldisulphide adducts of the individual monounsaturated PLFAs. Identi®cations were made using a Carlo Erba 5160 gas chromatograph
(column as above; He carrier gas, 10 psi head-pressure) coupled, using a heated transfer
line (320 8C), to a Finnigan MAT 4500 quadrupole mass spectrometer scanning in the
range of m/z 50 to 650 with a cycle time of 1.0 s (current was maintained at 300 mA with an
ion source temperature of 190 8C and an electron voltage of 70 eV). Stable carbon-isotope
compositions of individual compounds were determined using a Finnigan Delta S isotope
mass spectrometer coupled to a Varian 3400 gas chromatograph (column as above; He
carrier gas, 15 psi head-pressure) with an extensively modi®ed Finnigan type I combustion
interface. d13C values and associated errors were determined after correcting for the
isotopic composition of the exogenous methyl derivative group18,19. Column temperatures
were programmed from 40 8C (1 min isothermal) to 150 8C at a rate of 15 8C min-1, and
then to 240 8C at a rate of 4 8C min-1 with a ®nal isothermal period of 20 min.
Received 4 January 1999; accepted 18 February 2000.
1. Cicerone, R. J. & Oremland, R. S. Biogeochemical aspects of atmospheric methane. Glob. Biogeochem.
Cycles 2, 299±327 (1988).
2. Crutzen, P. J. Methane's sinks and sources. Nature 350, 380±381 (1991).
3. Hanson, R. S. & Hanson, T. E. Methanotrophic bacteria. Microbiol. Rev. 60, 439±471 (1996).
4. Conrad, R. Soil microorganisms as controllers of atmospheric trace gases (H2, CO, CH4, OCS, N2O,
and NO). Microbiol. Rev. 60, 609±643 (1996).
5. Boschker, H. T. S. et al. Direct linking of microbial populations to speci®c biogeochemical processes by
13
C-labelling of biomarkers. Nature 392, 801±804 (1998).
6. Hinrichs, K-U., Hayes, J. M., Sylva, S. P., Brewer, P. G. & Delong, E. F. Methane-consuming
archaebacteria in marine sediments. Nature 398, 802±805 (1999).
7. Zelles, L. & Bai, Q. Y., Rackwitz, R., Chadwick, & Beese, F. Determination of phospholipid and
lipopolysaccharide-derived fatty acids as an estimate of microbial biomass and community structures
in soils. Biol. Fertil. Soils 19, 115±123 (1995).
8. Bowman, J. P., Skerratt, J. H., Nichols, P. D. & Sly, L. I. Phospholipid fatty acid and liposaccharide fatty
acid signature lipids in methane utilizing bacteria. FEMS Microbiol. Ecol. 85, 15±22 (1991).
9. Nichols, P. D., Smith, G. A., Antworth, C. P., Hanson, R. S. & White, D. C. Phospholipid and
lipopolysaccharide normal and hydroxy fatty acids as potential signatures for methane-oxidizing
bacteria. FEMS Microbiol. Ecol. 31, 327±335 (1985).
10. Dun®eld, P. F., Werner, L., Henckel, T., Knowles, R. & Conrad, R. High af®nity methane oxidation by a
soil enrichment culture containing a type II methanotroph. Appl. Environ. Microbiol. 65, 1009±1014
(1999).
11. Nichols, P. D. et al. Detection of a microbial consortium, including type II methanotrophs, by use of
phospholipid fatty acids in an aerobic halogenated hydrocarbon-degrading soil column enriched with
natural gas. Environ. Toxicol. Chem 6, 89±97 (1987).
12. Guckert, J. B., Ringelberg, D. B., White, D. C., Hanson, R. S. & Bratina, B. J. Membrane fatty acids as
phenotypic markers in the polyphasic taxonomy of methylotrophs within the Proteobacteria. J. Gen.
Microbiol. 137, 2631±2641 (1991).
13. Bender, M. & Conrad, R. Effect of CH4 concentrations and soil conditions on the induction of CH4
oxidation activity. Soil Biol. Biochem. 27, 1517±1527 (1995).
14. Roslev, P. & Iversen, N. Radioactive ®ngerprinting of microorganisms that oxidize atmospheric
methane in different soils. Appl. Environ. Microbiol. 65, 4064±4070 (1999).
15. Hall, G. H., Simon, B. M. & Pickup, R. W. CH4 production in blanket bog peat: a procedure for
sampling, sectioning and incubating samples whilst maintaining anaerobic conditions. Soil Biol.
Biochem. 28, 9±15 (1996).
16. Bligh, E. G. & Dyer, W. J. A rapid method of total lipid extraction and puri®cation. Can. J. Biochem.
Physiol. 37, 911±917 (1959).
17. Guckert, J. B., Antworth, C. P., Nichols, P. D. & White, D. C. Phospholipid, ester-linked fatty-acid
pro®les as reproducible assays for changes in prokaryotic community structure of estuarine
sediments. FEMS Microbiol. Ecol. 31, 147±158 (1985).
18. Jones, D. M., Carter, J. F., Eglinton, G., Jumeau, E. J. & Fenwick, C. S. Determination of d13C values of
straight chain and cyclic alcohols by gas chromatography isotope ratio mass spectrometry. Biol. Mass
Spectrom. 20, 641±646 (1991).
© 2000 Macmillan Magazines Ltd
177
letters to nature
19. Rieley, G. Derivatization of organic compounds prior to gas chromatographic-combustion-isotope
ratio mass spectrometric analysis: Identi®cation of isotope fractionation processes. Analyst 119, 915±
919 (1994).
Acknowledgements
We thank J. Carter and A. Gledhill for their help with GC/MS and GCC/IRMS analyses and
C. Davies for setting up the long-term enrichment experiment. This study was supported
by the Natural Environment Research Council through the CEH Integrated Fund (N.R.P.,
P.I., G.H.H.) and through a Non-thematic Research Grant (P.I. and R.P.E.).
Correspondence and requests for materials should be addressed to R. P. E.
(e-mail: r.p.evershed@bris.ac.uk).
.................................................................
Infectious parthenogenesis
M. E. Huigens*, R. F. Luck², R. H. G. Klaassen*, M. F. P. M. Maas*,
M. J. T. N. Timmermans* & R. Stouthamer*
* Laboratory of Entomology, Department of Plant Sciences Wageningen
University, 6700 EH Wageningen, The Netherlands
² Department of Entomology, University of California, Riverside,
California 92521, USA
..............................................................................................................................................
Parthenogenesis-inducing Wolbachia bacteria are reproductive
parasites that cause infected female wasps to produce daughters
without mating1,2. This manipulation of the host's reproduction
enhances the transmission of Wolbachia to future generations
because the bacteria are passed on vertically only from mothers to
daughters. Males are dead ends for cytoplasmically inherited
bacteria: they do not pass them on to their offspring. Vertical
transmission of Wolbachia has been previously considered to
be the main mode of transmission. Here we report frequent
horizontal transmission from infected to uninfected wasp larvae
sharing a common food source. The transferred Wolbachia
are then vertically transmitted to the new host's offspring. This
natural and unexpectedly frequent horizontal transfer of parthenogensis-inducing Wolbachia intraspeci®cally has important
implications for the co-evolution of Wolbachia and their host.
Trichogramma kaykai (Hymenoptera; Trichogrammatidae) is a
minute egg parasitoid of a butter¯y species, Apodemia mormo
deserti (Lepidoptera; Lycaenidae), which inhabits the Mojave
Desert of southern California3. T. kaykai normally lays a clutch of
3±5 offspring in an A. mormo deserti egg. This parasitoid wasp
manifests haplo-diploid sex determination in which unfertilized
eggs develop into sons and fertilized eggs develop into daughters.
Between 6% and 26% of the T. kaykai females in these populations
are infected with parthenogenesis-inducing Wolbachia and produce
daughters from their unfertilized eggs4. These daughters are
produced when parthenogenesis-inducing Wolbachia suppresses
spindle formation during anaphase of the ®rst mitotic division,
thus restoring diploidy by fusion of the two mitotic nuclei5. In the
®eld, the offspring of two T. kaykai females can share and mature in
the same butter¯y egg6. Thus, we tested whether Wolbachia transfers
horizontally between the offspring of a Wolbachia-infected and an
uninfected T. kaykai female when they share the same egg and, if so,
whether this infection persists in its expression in subsequent
generations.
T. kaykai females collected at Last Chance Canyon, El Paso
Mountains, Kern County, California, were used to initiate infected
and uninfected lines. We cultured these lines in the laboratory on
eggs of the moth Trichoplusia ni 7, which do not harbour Wolbachia
(R. S., unpublished data). Each of these lines differed in the size of a
microsatellite DNA repeat. In our experiments, we offered a moth
egg to a female T. kaykai from one line, allowing her to lay a normal
clutch of 2±4 eggs. Two hours later, we exposed the same egg to a
178
female from the second line. In half the cases, a Wolbachia-infected
T. kaykai was offered the egg ®rst and in the other half the order was
reversed. We continuously observed and recorded the number of
eggs oviposited in a moth egg by each T. kaykai 8. The resulting
offspring were linked unambiguously to their parental female using
a microsatellite marker. If F1 offspring from a doubly parasitized egg
consisted of only females, we exposed these virgin F1 females
individually to host eggs and recorded the sex of their progeny.
Because only infected virgins produce daughters, their presence
in offspring of an F1 virgin originating from an uninfected
line indicated horizontal transfer of parthenogenesis-inducing
Wolbachia.
We found clear evidence for the horizontal transmission of
parthenogenesis-inducing Wolbachia and the expression of parthenogenesis. Twenty-one instances of horizontal transfer occurred
in 56 all-female broods from moth eggs that contained infected and
uninfected larvae (Table 1). Horizontal transfer to uninfected offspring occurred independent of the order in which the infected
and uninfected females were exposed to the host egg (x20.05,1 = 2.75;
n = 56). The bacteria's presence was con®rmed in all newly infected
females by means of polymerase chain reaction (PCR) using
Wolbachia-speci®c primers9. Both sons and daughters were produced by these newly infected, virgin F1 females, indicating inef®cient vertical transmission from mother to offspring. We suspect
that these females had insuf®cient Wolbachia titre to express
parthenogenesis in all of their offspring. To test whether a new
infection was maintained over several generations, we established
three lines, each of which consisted of offspring from a different,
newly infected female. Vertical transmission was inef®cient in the
®rst generation, but it was highly effective in the subsequent eight
generations. The transmission ef®ciency was 100% for 10 virgin
females of each line tested in generation 3 and generation 8.
The process by which uninfected Trichogramma larvae acquire
Wolbachia remains unclear. Several possible acquisition mechanisms exist. Infected larvae may die and be consumed by uninfected
larvae. Larvae may ®ght within the host egg and the uninfected
acquire an infection through blood-to-blood contact as found in the
woodlice Armadillidium vulgare10. Finally, an infected female may
inject Wolbachia into the host egg when she oviposits in it and the
larvae acquire the infection either orally or through wounds caused
by larval con¯ict. The ®rst possibility can be eliminated: in 3 out of
the 21 cases, all the eggs laid by the infected female survived to
become adults, yet some of the offspring from the uninfected female
became infected. We cannot distinguish between the two other
possibilities.
Our results show that offspring of uninfected females can acquire
suf®cient parthenogenesis-inducing Wolbachia to express parthenogenesis when they share a butter¯y egg with offspring of an
infected female. We know that such egg sharing occurs in the ®eld.
The maximum rates at which the offspring of Wolbachia-infected
and uninfected females share the same butter¯y egg in the ®eld can
be estimated from observed parasitism rates11. T. kaykai can
parasitize up to 80% of the Apodemia mormo deserti eggs, suggesting
that offspring of two or more females could share the same egg in as
many as 48% of cases. Up to 9% of the latter cases would involve
offspring of both infected and uninfected females (assuming a 10%
infection rate among females). Although such high parasitism rates
are infrequent, they illustrate the potential for horizontal transfer
under ®eld conditions.
Purely vertically inherited symbionts suffer from an accumulation of mutations similar to the Muller's ratchet in asexual
organisms12. However, horizontal transfer of parthenogenesisinducing Wolbachia among offspring of infected females offers the
potential for recombination between different Wolbachia variants
and thus for circumventing Muller's ratchet.
Vertical transmission has been considered the only common
mechanism by which parthenogenesis-inducing Wolbachia is trans-
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