diversity and salt tolerance of native acacia rhizobia isolated from saline and non-saline soils

Diversity and salt tolerance of native Acacia rhizobiaisolated from saline and non-saline soilsaec_1998 950..963

PETER H. THRALL,* LINDA M. BROADHURST, MOHAMED S. HOQUE ANDDAVID J. BAGNALLCSIRO Plant Industry, GPO Box 1600, Canberra ACT 2601, Australia (Email: [email protected])

Abstract Re-establishing native vegetation in stressed soils is of considerable importance in many parts of theworld, leading to significant interest in using plant–soil symbiont interactions to increase the cost-effectiveness oflarge-scale restoration. However, effective use of soil microbes in revegetation requires knowledge of how microbecommunities vary along environmental stress gradients, as well as how such variation relates to symbionteffectiveness. In Australia, shrubby legumes dominate many ecosystems where dryland salinity is a major issue, andimproving plant establishment in saline soils is a priority of regional management agencies. In this study, strains ofrhizobial bacteria were isolated from a range of Acacia spp. growing in saline and non-saline soils. Replicates of eachstrain were grown under several salinity levels in liquid culture and characterized for growth and salt tolerance.Genetic characterization of rhizobia showed considerable variation among strains, with salt tolerance and growthgenerally higher in rhizobial populations derived from more saline soils. These strains showed markedly differentgenetic profiles and generic affiliations to those from more temperate soils, suggesting community differentiation inrelation to salt stress. The identification of novel genomic species from saline soils suggests that the diversity ofrhizobia associated with Australian Acacia spp. is significantly greater than previously described. Overall, the abilityof some symbiotically effective strains to tolerate high salinity is promising with regard to improving host plantre-establishment in these soils.

Key words: legume, nitrogen-fixation, restoration, revegetation, salinity, soil symbiont.

INTRODUCTION

Ongoing loss and degradation of deep-rooted peren-nial vegetation is associated with desertification andsoil degradation, loss of biodiversity and decreasedproductivity of agricultural and grazing enterprises inmany parts of the world. Partly as a consequence ofsuch anthropogenic impacts there is an increasingfocus on the remediation of degraded landscapes andamelioration of environmental problems throughreforestation (Bell 1999; Broadhurst et al. 2008).Often, revegetation activities have focused on theinitial introduction of fast-growing woody pioneerspecies, many of which form associations with benefi-cial soil symbionts (Alnus spp. and Frankia (Monzon &Azcon 2001); Acacia spp. and rhizobial bacteria(Thrall et al. 2005); mycorrhizal fungi and a widerange of angiosperm hosts (Schwartz et al. 2006)).Thus, there is a growing recognition of the importanceof introducing appropriate beneficial soil symbionts toimprove plant establishment and growth, to aid in theremediation of degraded soils (Jasper 1994; Schwenke& Carú 2001; Rodríguez-Echeverría & Pérez-

Fernández 2005), or to improve the sustainability ofnative ecosystems (Hoquin et al. 2001).

Re-establishing effective plant–symbiont interac-tions is of particular importance in degraded or low-fertility soils, or where native soil communities havebeen altered as a result of long-term cultivation orcropping (Zahran 1999; Requena et al. 2001; Thrallet al. 2001, 2005; McKinley et al. 2005). For example,mycorrhizal fungi are being used to increase establish-ment of vegetation in desertified phosphorus-deficientmediterranean and other degraded ecosystems(Herrera et al. 1993; Cuenca et al. 1998, 2002; Cara-vaca et al. 2004; Medina et al. 2004), or in bioreme-diation of mine-sites (Perotto & Martino 2001).Nitrogen-fixing Frankia spp. and actinorhizal hostplants have also been utilized to improve plant perfor-mance in similar situations (Schwenke & Carú 2001;Markham 2005). Similarly, woody legumes and theirassociated rhizobial symbionts are key components oflarge-scale replanting efforts aimed at restoringnitrogen-depleted soils in a diversity of ecosystemsranging from semi-arid African savannas (Diabateet al. 2005) and dry mediterranean habitats (Requenaet al. 1997) to subtropical and tropical forests in Chinaand South America (Franco & de Faria 1997; Penget al. 2005) and native tall-grass prairies in NorthAmerica (Tlusty et al. 2004).

*Corresponding author.Accepted for publication December 2008.

Austral Ecology (2009) 34, 950–963

© 2009 The Authors doi:10.1111/j.1442-9993.2009.01998.xJournal compilation © 2009 Ecological Society of Australia

In Australia, there is a high diversity and abundanceof Acacia spp. and other native shrubby legumes(Groves 1994; Maslin & MacDonald 2004). Acaciaspp. also possess varying degrees of adaptation tosaline soils (Aswathappa et al. 1986; Marcar & Craw-ford 2004), and have considerable promise for thereclamation of salt- or acid-affected land (Ansari et al.1998; Zahran 1999; Rodríguez-Echeverría & Pérez-Fernández 2005). This is particularly relevant in theAustralian context, where establishment of deep-rooted perennial vegetation is viewed as an essentialcomponent of protecting native biodiversity, restoringwater balances and reducing the spread of drylandsalinity (Hobbs et al. 1993; Barrett-Lennard 2002;Peck & Hatton 2003).

Results from large-scale direct-seeding field trialshave shown that host-plant establishment, growth andsurvival can be significantly improved when inoculatedwith appropriate native strains of Bradyrhizobium(Thrall et al. 2005). Several previous studies haveevaluated the salt-tolerance of native rhizobia with theaim of using these in bioremediation of saline soils (Lal& Khanna 1994; Arun & Sridhar 2005). Acacia rhizo-bia differ in their ability to tolerate salinity (Zhanget al. 1991; Lal & Khanna 1994; Surange et al. 1997;Hashem et al. 1998); some strains are able to survive atsalt concentrations up to 800 mM (Merabet et al.2006), while others show increased growth rates undersaline conditions relative to controls (Jenkins 2003),implying some degree of evolutionary adaptation tosaline soils. In some cases, the nitrogen-fixing ability ofsalt-tolerant strains decreases by up to 75% withincreasing salinity (Lal & Khanna 1994). However,other strains have been shown to maintain their sym-biotic capacity up to 200 mM NaCl (Zou et al. 1995).Craig et al. (1991) found that salt-tolerant strains ofAcacia rhizobia in combination with a highly salt-tolerant provenance of A. redolens formed associationsthat did not differ in the level of benefits conferredirrespective of salt concentration up to 160 mM. Incontrast, on a less salt-tolerant provenance of A. redo-lens, and on A. cyclops, both infectivity and N2-fixingeffectiveness decreased as the external salt concentra-tion increased.

Despite the ecological importance of native legumesin many natural ecosystems, the diversity and distri-bution of associated root-nodule bacteria remainspoorly understood. Lafay and Burdon (1998) isolatedrhizobia from a diversity of shrubby legume speciesoccurring naturally across temperate south-easternAustralia, and characterized these into genomicspecies using restriction fragment length polymor-phism (RFLP) molecular markers of the 16S smallsubunit (SSU). Most of the 745 isolates (94.5%) wereclassified into 16 genomic species within Bradyrhizo-bium, with the few remaining isolates identified asbelonging to Rhizobium and Mesorhizobium. A similar

study of isolates from Acacia spp. in north WesternAustralia found both Rhizobium and Bradyrhizobiumpresent (Yates et al. 2004). Other studies suggest thatrhizobial community structure may vary along envi-ronmental stress gradients (Barnet & Catt 1991;Thrall et al. 2008).

The current study is part of a larger research pro-gramme aimed at improving the establishment ofnative woody perennial plants in saline dischargezones. A primary goal was to assess the extent of varia-tion for salt tolerance in native populations of Acaciarhizobia. However, given the results from previousstudies, which suggest that the rhizobia nodulatingnative shrubby legumes in stressed soils may differfrom those occurring in non-stressed situations(Barnet & Catt 1991), we also genetically character-ized a subset of strains from saline and non-saline sitesand compared these to known genomic species.Strains from non-saline soils were derived from abroad range of Acacia species native to southeasternAustralia (Lafay & Burdon 1998). Strains from salinesoils were isolated from the rhizosphere of several salt-tolerant Acacia species growing in sites with varyinglevels of soil salinity.

METHODS

Rhizobial isolates from non-saline soils

A group of 85 rhizobial isolates were selected from anexisting CSIRO Plant Industry collection on the basisof their ability to promote host plant growth, as estab-lished in previous glasshouse trials (Burdon et al. 1999;Thrall et al. 2000; Murray et al. 2001). These isolateswere ranked in the highest 10% of those tested withregard to N2-fixing effectiveness and were thus consid-ered to represent ‘elite’ strains (hereafter referred to asGroup E), and therefore likely to be desirable for res-toration applications. The locations and host speciesfrom which these isolates were originally collected aregiven in Table 1. The Acacia spp. from which theseisolates were originally sampled are known to onlyexhibit low to slight tolerance to saline soils (Marcar &Crawford 2004). None of the collection sites werelocated in areas known to be saline (Soil ConservationService of NSW 1989; Doyle & Habraken 1993).

Rhizobial isolates from saline soils

In October of 2004, soil samples were collected fromthe rhizosphere of natural stands of several Acaciaspecies known to possess at least some degree of salttolerance (Marcar & Crawford 2004) from sites inVictoria and New SouthWales (Table 2). For each site,

SALT TOLERANCE OF ACACIA RHIZOBIA 951


5–10 soil samples were taken haphazardly across thesite to a depth of 100 mm and then bulked. Soil salin-ity was measured as electrical conductivity of a 1:5soil : water extract with appropriate soil texture con-version factors, and soil pH was measured as pHCa

(0.01 M CaCl2) (Marcar & Crawford 2004).

To isolate rhizobia from the soil samples, 250-mmdiameter pots containing 1:1 sterilized vermiculite :sand mixture were used (five replicate pots per bulkedsoil sample). For each pot 100 g of a given soil wasplaced over this mixture and covered with a secondlayer of vermiculite : sand. For each soil, seedlings ofthe host Acacia species growing at that site were usedto trap rhizobia (3–5 seedlings/pot). Acacia seed(obtained from the Australian Seed Company, PO Box67, Hazelbrook NSW 2779) was pretreated withboiling water for 1 min, allowed to cool, surface ster-ilized with ethanol (30 s), followed by 5% (v/v) sodiumhypochlorite (10 min) and rinsed with distilled water.Plants were grown under standard glasshouse condi-tions, watered with N-free 1 : 20 diluted McKnight’ssolution (McKnight 1949) three times weekly, andUV-sterilized tap water otherwise. Plants were har-vested after 2–5 months. Nodules were individuallyexcised from roots, soaked in distilled water for 2 h,surface sterilized, crushed and streaked onto yeast-extract mannitol agar (Vincent 1970) plates and incu-bated at 28oC for 5–10 days. Of the 755 isolatesobtained, a subset of 86 (hereafter referred to asGroup S) were haphazardly chosen for the salt toler-ance experiments.

Growth rate and salinity responses

A total of 85 Group E isolates (elite strains from non-saline soils) and 86 Group S isolates (those from salt-tolerant acacias growing in saline soils) were tested forsalt tolerance in liquid culture. Treatments included200 mM, 400 mM and 800 mM NaCl concentration(saline soils in Australia are dominated by sodiumchloride, sodium bicarbonate and sodium carbonate;Rengasamy (2002), as well as a salt-free control. Iso-lates were sub-cultured, then grown on slants of yeastmannitol agar (YMA) for one week and sub-culturedonto a second series of YMA slopes for a further4–5 days.Transfers from slants were used to inoculate50 mL of yeast extract mannitol broth. Each broth wasincubated at 120 rev min-1 and 28°C in a ThermolineTSIR-406-25 orbital shaking incubator. For eachsalinity treatment, the growth of three replicates foreach isolate was measured daily for 3 days. Althoughmore frequent measurements would have alloweddetailed characterization of individual growth curves,in contrast to many previous studies which have onlyexamined limited isolate numbers, this approachenabled us to undertake a broad survey of variation insalt tolerance across a diverse collection of rhizobia.Due to space constraints the entire set of 171 isolatescould not be simultaneously grown. Accordingly, iso-lates were evaluated in several consecutive batcheswith a control isolate (CPI51) regrown in each run asa control to assess potential differences in experimen-

Table 1. Host species and site locations for Group E iso-lates from the CSIRO Plant Industry collection

Host species Isolate # and collection sites

A. binervata 7203, 7222, 7223, 7225 (Kiama, NSW)A. binervia 7132, 7133, 7137 (Richmond NSW)A. blayana 6421 (Wadbilliga NP, NSW)A. cangaiensis 9B-19 (Cangai, NSW)A. cincinnata 50B9 (Gympie, QLD)A. dangarensis 1406, 1407 (Mt. Dangar, NSW)A. dealbata 4506 (Inglis River, TAS); 1604

(Kandos, NSW); 3301, 3323, 3335,3337 (Ben Lomond, NSW); 46A9,46B13 (Mt. Elephant, TAS); 47A17,47A21,47B13 (Snug, TAS)

A. deanei 52A23, 52B20 (E. Goodawindi, NSW);6921, 6925, 6935 (W. Wyalong,NSW)

A. decurrens 7506, 7525A, 7525B (N. Goulburn,NSW); 7604 (Picton, NSW)

A. elata 7422B, 7420B (Buxton, NSW); 10A12,10B16, 10B21 (Gloucester, NSW)

A. filicifolia 7010, 7037 (Yadboro Flat, NSW)A. glaucocarpa 51B20, 51B21 (Gayndah, QLD)A. implexa 402, 409, 433 (Moonan Flat, NSW);

2235 (Glenmaggie Lake, VIC); 2527(Pyalong, VIC); 53B9, 53B13(Sofala, NSW)

A. irrorata 706 (Wauchope, NSW); 6721, 6722(Bodalla, NSW)

A. leucoclada 48A15, 48A20, 48B16, 48B22, 48B25(Inverell, NSW)

A. mearnsii 1710 (Cooma, NSW); 2604 (NEKyneton, VIC); 3607 (Berrima,NSW)

A. melanoxylon 1236 (Singleton, NSW); 2011 (GoannaCreek, VIC); 2913, 2924 (Mt.Gambier, SA); 2836 (GellibrandRiver, VIC); 3910 (Lileah, TAS);49A21, 49B1, 49B5, 49B13(Cascade, NSW); 3301, 3323, 3335,3337 (Bli Bli, QLD); 7723A(Gellibrand SF, VIC)

A. parramattensis 6615A, 6625B (Numeralla, NSW)A. nanodealbata 2706, 27A22 (Laver’s Hill, VIC)A. parvipinnula O310, 3A21 (Howes Valley, NSW)A. rubida 8007, 8013 (Queanbeyan, NSW)A. silvestris 55A8, 55A10, 55B9 (Narooma, NSW)A. trachyphloia 54A16 (Monga SF, NSW); 6319B

(Bateman’s Bay, NSW)

NP, National Park; NSW, New SouthWales; QLD, Queen-sland; SA, South Australia; SF, State Forest;TAS,Tasmania;VIC, Victoria.

952 P. H. THRALL ET AL.

© 2009 The Authorsdoi:10.1111/j.1442-9993.2009.01998.xJournal compilation © 2009 Ecological Society of Australia

tal conditions. This isolate showed near-identicalresponses across experimental runs (data not shown).

Isolate growth rates were measured spectrophoto-metrically at 660 nm (Jenkins 2003) using a BeckmanDU-50 spectrophotometer to assess changes inturbidity. Specific growth rates (m) were calculatedfrom the natural log of the OD660 readings against timefor the linear portion of the growth curves (i.e. theslope over the first 24 h of exponential growth (Zwi-etering et al. 1990; Jenkins 2003)). Responses to salin-ity were assessed as the percentage difference betweenthe 3 day OD660 reading at a given concentration ofNaCl and the 3 day reading for the salt-free control.This index of salt response was used to rank isolateperformance for each of the salt concentrations.

Genetic characterization of isolates

A selection of 130 bacterial isolates from both saline(n = 47) and non-saline (n = 83) soils were chosen forgenetic characterization. Bacterial DNA was preparedfollowing the methods described by Lafay and Burdon(1998). Single colonies collected from bacteria grownon YMA plates were suspended in 100 mL of 10 mMTris (pH 8.0)–1 mM EDTA–1% Triton X-100 solu-tion and boiled for 5 min. The solutions were brieflycentrifuged, washed with 100 mL of chloroform andthe supernatant used as polymerase chain reaction(PCR) templates. Twenty-eight isolates representing12 genomic species previously characterized by Lafayand Burdon (1998) were similarly treated for compari-son with the study isolates. These included strains ofBradyrhizobium (A, B, F, I, J, K, M, O, P), Mesorhizo-bium (S, T) and Rhizobium (Q).

The 16S f27 forward and 16S r1392 reverse primers(Lane 1991) were used to amplify the SSU. PCR wascarried out in 100-mL reactions containing 10 mL of

template DNA (10 hg mL-1), 10X PCR buffer (PerkinElmer), 5 pM of each primer (Sigma), 5 mM of eachdNTP (Invitrogen), 25 mM MgCl2 (Perkin Elmer),Taq (Perkin Elmer, 5 U mL-1). Amplification was per-formed under conditions specified by Lafay andBurdon (1998) with the exception that an annealingtemperature of 55°C was used. A 20-mL aliquot of thisPCR product was digested with one of four restrictionendonucleases (HhaI, HinfI, MspI and RsaI; NewEngland Biolabs) according to the manufacturersguidelines which had previously been found to dis-criminate among strains at the level of genomic species(Lafay & Burdon 1998). Digested products were sepa-rated on a 2% NuSieve agarose gel run at 180 V for2 h, visualized with ethidium bromide and thenphotographed. Restriction profiles based on the fourenzymes were determined for each isolate and com-pared with those of the known genospecies. Isolateprofiles with 100% banding pattern matches for allfour enzymes were considered to be identical whileisolates with a difference in one or more of the fourenzymes were considered to represent new genomicprofiles. Each of these was consecutively numbered.The PCR-RFLP profiles of each genomic species wereused to construct a binary matrix (1/0) indicating thepresence/absence of bands. This data were used togenerate a similarity matrix based on Euclidean dis-tances and principal co-ordinates calculated for thegenomic species. These co-ordinates were then visual-ized in 2-dimensional space.

To determine isolate generic affiliations, representa-tives of each genospecies were sequenced followingamplification using the 16S f27 and 16S r1392primers. Bands were excised from agarose gels,cleaned with the UltraClean(tm) DNA PurificationKit (Mo Bio Laboratories) as specified by the manu-facturer and used as template for separate sequencingreactions using the forward (16S F27) and reverse

Table 2. Host species and site location for Group S isolates derived from saline soils in Victoria and New South Wales

Host species Isolate # and collection sites Soil typeConductivity

(dS m-1) pHCa

A. montana CPI250, CPI402–407, CPI509, CPI536-CPI538, CPI542(Wills Bend, VIC)

Medium clay 2.4 6.5

A. pendula CPI252, CPI267, CPI506, CPI510, CPI513, CPI543–549,CPI551–564 (Coleambally, NSW)

Medium clay 12.6 7.4

A. salicina CPI248–249, CPI261–264, CPI311, CPI313, CPI512,CPI521–527, CPI539–540, CPI565–568 (Patho, VIC)

Medium clay 67.5 7.6

A. sophorae CPI246–247, CPI314, CPI530–535, CPI541, CPI572–574(Broulee, NSW)

Sand 3.0 7.7

A. stenophylla CPI243–244, CPI251, CPI265–266, CPI312, CPI408, CPI505,CPI511, CPI520, CPI528, CPI529 (Mystic Park, VIC)

Light clay 10.4 7.6

Salt tolerance of Acacia pendula, A. salicina and A. stenophylla is described in Marcar and Crawford (2004). A sophorae is acoastal foredune species able to tolerate full exposure to salt spray; salt tolerance has not been assessed for A. montana, althoughthe Australian National Botanical Gardens website lists this species as suitable for coastal plantings, and anecdotal reportssuggest that it can occur in moderately saline sites.



(16S R1392) primers with ABI Big Dye terminatoraccording to the manufacturers protocols. Reactionswere then analysed on an ABI 3730 capillarysequencer. Forward and reverse sequences for eachisolate were manually aligned to check for sequencequality using MEGA4 (Tamura et al. 2007) and theforward sequences used for a gene bank BLASTsearch to help identify isolates. A subset of thesesequences identified isolates belonging to non-nodulating bacterial groups previously shown to occuras endosymbionts in rhizobial nodules (e.g. Zakhiaet al. 2006; Muresu et al. 2008). These isolates werere-inoculated onto several Acacia species to confirmtheir non-nodulating status.

Statistical analyses

One-way analyses of variance were used to assess dif-ferences between the Group E and Group S isolateswith regard to specific growth rate (m), and toleranceto salt (defined as % change in maximum OD660 for the3-day readings). Repeated measures ANOVAs wereperformed to further examine how rhizobial growthand salt tolerance changed across salinity levels for thetwo isolate groups. Analysis of variance was also usedto examine among-site differences in salt tolerance forthe Group S isolates (this analysis was precludedfor Group E because of the low number of isolatesper site). Pearson product–moment correlations wereused to evaluate whether the degree of salt tolerance(% change in maximum density) shown by isolates wasconsistent for different treatments, and to examinethe relationships between specific growth rate (m),maximum OD660 and salt-tolerance. Kendall’s coeffi-cient of concordance (W) was calculated to examinethe degree to which rank order for growth rate andsalt-tolerance was preserved across different salt con-centrations for individual isolates.

RESULTS

Variation in rhizobial growth rates

Initial bacterial growth rates varied more than three-fold from slowest (m = 0.027 for a Group E strainisolated from Acacia nanodealbata) to fastest (m =0.147 for a group S isolate from A. montana) over thefirst 24 h for the 0 mM NaCl control treatment.Whilethe distribution of growth rates for Group S andGroup E strains overlapped considerably (Fig. 1), asshown by one-way analyses of variance, the Group Srhizobia were on average significantly faster growingthan the Group E isolates from non-saline sites. Thiswas particularly the case for the 0 mM NaCl (mean

growth rates: 0.071 and 0.085 for Group E and GroupS respectively; F1,169 = 23.25, P < 0.0001), 200 mMNaCl (mean growth rates: 0.061 and 0.079;F1,169 = 37.53, P < 0.0001) and 400 mM NaCl (meangrowth rates: 0.059 and 0.066; F1,169 = 6.05, P =0.015) treatments.

As confirmed by a repeated measures anova, notonly were there significant overall differences ingrowth rates across salinity levels (F3,507 = 213.14,P < 0.0001) and between Group E and Group S iso-lates (F1,169 = 13.45, P < 0.001), but there was a highlysignificant interaction between isolate group and salin-ity level (F3,507 = 20.01, P < 0.0001).The largest differ-ences between the two groups was seen for 200 mMNaCl as some of the more salt-tolerant Group S iso-lates actually grew faster at this concentration than at0 mM NaCl. Clearly however, with increasing saltconcentrations beyond 200 mM NaCl, the differencesin distribution of growth rates between the two groupsdiminished, and in fact, at 800 mM NaCl, averageinitial growth rate did not differ between the Group Sand Group E isolates (0.049 for both groups). Thisreflected the fact that at very high salt concentrations,very few isolates were able to persist.

Variation in salt-tolerance of rhizobial isolates

There was a broad diversity of responses of the 171rhizobial isolates to salt (Fig. 2). For example, at aconcentration of 200 mM NaCl, the percentage differ-ence in the 3 day OD660 readings relative to thatobserved in the zero salt control varied from -71.6%(an extremely salt-sensitive Group E isolate from A.implexa) to +57.9% (a highly salt-tolerant Group Sisolate from A. sophorae). At this concentration, 26.7%of the Group S isolates showed increased densities byday 3 relative to the zero salt controls. This was incontrast to the Group E isolates where only 5.9% of thestrains performed better at 200 mM NaCl than in theno-salt controls (similar results were seen for the400 mM vs. control comparisons, 17.4% vs. 3.5% forGroup S and Group E isolates, respectively). The factthat at least some isolates, particularly those from salinesoils, actually showed higher final densities in 200 mMand 400 mM solutions than in the zero salt controls wassuggestive of significant levels of adaptation to salinesoils. However, it is worth noting that both salt-sensitiveand salt-tolerant strains were present within sites,showing that even in saline soils there could be signifi-cant variation in tolerance (e.g. at 200 mM NaCl, onestrain isolated from A.pendula at Patho showed a 63.9%decline in maximum density from the zero control,while another showed a 47.6% increase).

Similar to the results for initial growth rates, one-wayanalyses of variance showed that the Group S rhizobiawere on average significantly more salt-tolerant than



the Group E isolates from non-saline sites. Thus, themean % difference in 3-day OD660 readings between the200 mM and zero salt treatments was -33.5% and-10.4% for the Group E and Group S isolates(F1,169 = 45.10, P < 0.0001).While still significant, dif-ferences in salt tolerance decreased for the 400 mMcomparison (mean % change in density from 0 NaCl:-38.1% and -26.5% for the Group E and Group Sisolates, respectively; F1,169 = 10.26, P = 0.002). Therewas no difference between isolate groups at 800 mMNaCl (mean % change in density: -55.7% and -57.7%for Group E and Group S; F1,169 = 0.69, P = 0.41).Therepeated measures anova demonstrated major effects ofrhizobial group (F1,169 = 14.62, P < 0.001) and salinitylevel (F2,338 = 387.19, P < 0.0001) on salt tolerance, aswell as a significant interaction between these variables(F2,338 = 48.33, P < 0.0001), confirming that salt toler-ance of the two rhizobial groups varied across salinitylevels.

To assess whether the ability of individual isolates totolerate salt varied with different concentrations,isolates within each group were ranked with regard tosalt tolerance (as defined above) for each of the threesaline treatments (200, 400 and 800 mM NaCl) andKendall’s coefficient of concordance (W) wascalculated.The results showed that for both the GroupE and Group S isolates, rank order did not changesignificantly across salt treatments (W = 0.889,c2 = 224.0, P < 0.001 for Group E; W = 0.752,c2 = 189.6, P < 0.001 for Group S) indicating that thedegree of salt tolerance was largely independent ofparticular salt concentrations.The actual magnitude ofsalt tolerance (% change in 3-day OD660 readings) wasalso positively correlated for the 200 versus 400 mMNaCl treatments (Group E: r = 0.86, P < 0.0001;Group S: r = 0.76, P < 0.0001) and 200 versus800 mM NaCl (Group E: r = 0.78, P < 0.0001; GroupS: r = 0.52, P < 0.0001).

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Fig. 1. Distribution of specific growth rates (m) for the 85 Group E isolates (derived from non-salt-tolerant Acacia speciesgrowing in presumed non-saline soils; Table 1) and the 86 Group S isolates (isolated from a range of sites dominated bysalt-tolerant Acacia spp.; Table 2). Gray bars represent the Group E isolates, and open hatched bars represent the Group Sisolates (areas of overlap between distributions are shaded as well as hatched): (a) control treatment (0 mM NaCl); (b) 200 mMNaCl; (c) 400 mM NaCl; and (d) 800 mM NaCl. The x-axis represents salt-tolerance, with large negative values equating tohighly sensitive strains and large positive values representing highly tolerant strains. The y-axis represents the proportion ofisolates in different tolerance categories.



Overall, these results provide additional evidencethat faster growth may be linked to the ability to tol-erate stressful environments, although it should benoted that faster growth per se, does not necessarilyconfer a greater degree of salt tolerance as a number offast-growing isolates were also shown to be salt-sensitive. Although the trend was for faster-growing,more salt-tolerant bacteria to be found in sites withhigher levels of soil salinity (e.g. compare Fig. 3a,d),there were a number of examples where salt-tolerantstrains were found in non-saline soils. For example,some slower growing strains isolated from A. decurrensat the non-saline North Goulburn site showed positivegrowth in salt solutions up to 400 mM NaCl (Fig. 3c).The converse was also observed – at the highly salinePatho site strains were recovered that grew poorly evenin low to moderate salt solutions (Fig. 3b). Overall, forthe Group S isolates, despite the generally higher levelsof salt tolerance, there was considerable within-sitevariability in salinity responses and results from one-

way ANOVAs showed that there were no consistentdifferences among the Group S sites in either growthor salt-tolerance. It is also worth noting that the rhizo-bial population present at the most saline site (Patho,A. salicina; Table 2) did not generally have the highestaverage tolerance, although the only Group S strainsable to grow at 800 mM NaCl were derived from thissite.

Characterization of rhizobial isolates

Each enzyme produced a different number of restric-tion banding patterns across the 130 isolates – HhaIproduced 12 patterns, HinfI produced 19, MspI pro-duced 21 and RsaI produced 15. Despite the largenumber of banding patterns produced by the fourenzymes, genomic species ‘A’, ‘F’ and ‘M’ previouslycharacterized by Lafay and Burdon (1998) from sitesin temperate southeastern Australia, could not be dis-

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Fig. 2. Distribution of salt tolerance (defined as % change in max OD660 readings at a given salt level relative to the no saltcontrols) for the Group E (gray bars) and Group S (open hatched bars) isolates (areas of overlap between distributions areshaded as well as hatched): (a) values for the 200 mM NaCl treatment; (b) 400 mM NaCl treatment; and (c) 800 mM NaCltreatment. The x-axis represents salt-tolerance, with large negative values equating to highly sensitive strains and large positivevalues representing highly tolerant strains. The y-axis represents the proportion of isolates in different tolerance categories.



tinguished from each other and isolates with thisrestriction pattern were pooled into a single group(AFM). In addition, isolates previously characterizedas genomic species ‘I’ had two banding patterns, one ofwhich was consistent with genomic species ‘J’ andthese indistinguishable isolates were also pooled (‘IJ’)with the remaining isolates designated as genomicspecies ‘I’.

In total, 28 PCR-RFLP profiles were identifiedacross the 130 isolates. Of these, only profiles I, IJ,AFM & 11 were shared by Groups E and S (Fig. 4). InGroup E 97% of the isolates had restriction patternsidentical to a subset of the known genomic speciesincluded in the study with most belonging to theknown bradyrhizobial groups ‘AFM’ (50%), ‘B’(26%), ‘I’ (12%) and ‘IJ’ and ‘P’ (both 4%). Of theremaining isolates, sequence data confirmed one ofthese as Rhizobium (profile 15, Table 3), one asbelonging to the nodulating genus Herbaspirillum(profile 11), and a third as a non-nodulating Mycobac-

terium sp. (profile 32, Table 3). The non-nodulatingstatus of this isolate was confirmed by re-inoculationonto multiple Acacia seedlings (Table 3). In contrast,only 17% of the Group S isolates were from knowngenomic species (‘AFM’, 4%; ‘IJ’, 11%; ‘I’, 2%) withthe remaining 83% of isolates being represented by 20new profiles. Twelve of these profiles (10, 18–21, 24,26–31; 40% of Group S) were identified by thesequence data as being non-nodulating bacteria whichwas also confirmed by re-inoculation (Table 3). Of theremaining isolates, profile 16 (Sinorhizobium) was themost abundant (21%) with the other isolates rangingfrom 2% to 4% of the total soil profile. Sequence data(Table 3) indicated that the remaining isolatesbelonged to Bradyrhizobium (profile 1), Rhizobium(profiles 12, 13, 19), Mesorhizobium (profile 32) andSinorhizobium (profile 17).

Principal co-ordinates analysis of the genomicspecies identified by PCR-RFLP fragment patternsdivided the isolates into several major groups (Fig. 5)

0.0

0.5

1.0

1.5

a

0.0

0.5

1.0

1.5

0.0

0.5

1.0

1.5

0.0

0.5

1.0

1.5

Days in culture

Op

tica

l d

ensi

ty (

660n

m)

b

dc

0 1 2 3 0.1 1.2 2.3

0 1 2 3 0.1 1.2 2.3

Fig. 3. Representative responses of salt-tolerant and salt-sensitive isolates from saline and non-saline soils to 0 (filled circles),200 (open triangles), 400 (filled squares) and 800 (open circles) mM NaCl salt solutions: (a) a salt-tolerant strain (CPI54) fromAcacia salicina growing at Patho, Victoria (a highly saline site); (b) a salt-sensitive strain (CPI49) from Patho; (c) a salt-tolerantstrain (7525A) from A. decurrens growing at a non-saline site near North Goulburn, New South Wales; and (d) a salt-sensitivestrain (7506) from the North Goulburn site. Standard error bars are shown.



with the first two principal co-ordinates accounting forapproximately 32% of the variation present. The pre-viously identified Bradyrhizobium genomic species(AFM, B, I, IJ, K, O and P) were separated intopositive PCord1 space whereas Rhizobium genospeciesQ was in negative PCord1 and PCord2 space.The twoMesorhizobium isolates (S and T) fell between thesetwo groups although closer to the Bradyrhizobiumgroup. Several of the newly identified isolate profileswere either identical to, or closely affiliated with knowngenospecies of Bradyrhizobium. For example, profiles4 and 7 were identical to ‘I’ and ‘AFM’ respectively,whereas profile 2 was associated with this bradyrhizo-bial group but did not match any known genospecies.No isolate profile was identical to Rhizobium genospe-cies Q but several isolates were closely associated,including the two sinorhizobial isolates, 16 and 17. Asingle isolate (profile 32) was affiliated with the twoMesorhizobium genospecies S and T. Many of the newprofiles that were confirmed to be non-nodulating bac-teria were distributed broadly in negative PCord1space with several grouping with the nodulating genusHerbaspirillum.

DISCUSSION

Rhizobial growth and salt tolerance

A number of previous studies have suggested that fastgrowing isolates are generally more salt-tolerant thanslow growing isolates (Graham & Parker 1964; Balaet al. 1990; Marsudi et al. 1999; Mohamed et al. 2000;Yates et al. 2004). Barnet and Catt (1991) character-

ized the growth rates of Acacia rhizobia from bothtemperate and arid sites. Fast growing isolates wereexclusively present at the arid Fowler’s Gap siteleading the authors to speculate that these isolatesgrouped with Rhizobium, while the slow growingstrains from a more temperate site were likely tobelong to Bradyrhizobium. Similar studies ofWest Aus-tralian native legumes (including Acacia spp.) haveshown that fast-growing Rhizobium were able tosurvive and grow in 170 mM NaCl and also at 37°C,whereas slow-growing Bradyrhizobium were less viableunder these conditions (Marsudi et al. 1999; Yateset al. 2004). In our study, isolates from saline sitestended to be faster growing (Fig. 1) and on average,more salt-tolerant (Fig. 2), but faster growth was notcompletely correlated with salt tolerance as somerapidly growing isolates were also salt-sensitive. More-over, there were clear exceptions to the traditionallyheld view that fast-growing salt-tolerant isolates arealways Rhizobium rather than Bradyrhizobium. Thus,some fast-growing salt-tolerant strains (e.g. strain50B9 from Acacia cincinnata; Table 1) had been char-acterized as Bradyrhizobium in previous studies (Lafay& Burdon 1998), while other strains of Bradyrhizobiumthat were relatively slow growing were also salt-tolerant (e.g. strain O310 from A. parvipennula;Table 1).

Patterns of variation in genomic species inrelation to soil salinity

Of considerable interest was the difference in rhizobialcommunity structure between the two soil classes. For

Fig. 4. Distribution of nodule isolates into genomic species based on PCR-RFLP banding patterns. Previously characterizedgenomic species are designated by capital letters according to Lafay and Burdon (1998, 2001), and those belonging to novelgenomic species are labeled with numbers: (a) Group E isolates from non-saline sites; and (b) Group S isolates from saline sites.Grey shading indicates isolates found in one soil type only, dappled shading indicates isolates found in both soil types, and blackshading indicates non-nodulating endosymbiotic bacteria.



example, 96% of the isolates from the non-saline soilsgrouped with Bradyrhizobium, as compared with only19% of those from saline soils. In contrast, 34% ofisolates from saline soil were Rhizobium, comparedwith 1% in non-saline soils suggesting major rhizobialcommunity shifts in relation to soil salinity levels. Theearlier studies of Lafay and Burdon involved samplingrhizobial communities from temperate sites in NewSouthWales, Queensland,Tasmania andVictoria char-acterized by higher rainfall and most likely lower salin-ity levels (National Land and Water Resources Audit2001a) than the saline sites represented by the GroupS isolates in this study. While our isolates were col-lected from the rhizospheres of Acacia spp. notsampled by Lafay and Burdon (1998, 2001), it isworth noting that their studies included not only anumber of acacias, but also a broad range of othershrubby legume hosts (e.g. Daviesia, Pultenaea,Bossiaea). It therefore seems likely that the range ofnew genomic species identified in the current study isa consequence of sampling more saline soils, ratherthan being due to the identity of particular hostspecies.

Contrasting patterns in isolate abundance were alsoobserved between the two soil types. Approximately25% of the non-saline isolates belonged to genospeciesB (profile 8) and although this genospecies has beenisolated from eight other shrubby legume taxa occur-ring in southeastern Australia, it was absent from thesaline soil isolates. Overall, profile 7 (‘AFM’) was themost abundant non-saline profile (50%). While thisgenospecies appears to be widely distributed, havingbeen isolated from 17 other legume taxa (Lafay &Burdon 1998, 2001) it represented only a small pro-portion (4%) of isolates from saline soils. Of furtherinterest was the high number of non-nodulating endo-symbiotic isolates (approx. 40%) identified from salinesoils. There is evidence that a diverse array of endo-phytic bacteria coexist within rhizobial nodules(Zakhia et al. 2006; Muresu et al. 2008), and thatthese may, in some cases, play a role in plant growthpromotion or in the provision of other benefits (e.g.enhancing nodulation, protection against disease).Further research is clearly required to assess the func-tional relationships between these non-nodulatingbacteria, rhizobia and their host plants, particularlyalong environmental stress gradients.

Salt tolerance and revegetation ofsalinized landscapes

Dryland salinity has led to loss of agricultural produc-tivity and biodiversity, and destruction of infra-structure which has threatened the survival of farmingcommunities and towns across significant areas inAustralia (National Land and Water Resources Audit

Table 3. Most likely generic affiliations of isolates using16S sequence data following a Genebank BLAST search

IsolateprofileNo.

Generic affiliation genebank ID(sequence match percentage)

1 Bradyrhizobium japonicum EU145982 (99%);2 Bradyrhizobium betae AY372184 (99%);

Bradyrhizobium japonicum DQ786800 (99%)4 Bradyrhizobium japonicum DQ786800 (99%)5 Bradyrhizobium japonicum AF363126 (99%);

Bradyrhizobium canariense AY577427 (99%)7 Bradyrhizobium japonicum AB195988 (99%)8 Bradyrhizobium japonicum AY904765 (99%)

10 Curvibacter gracilis AB109889 (100%);Pseudomonas lanceolata AB021390 (99%);Aquamonas fontana AB120966 (98%)

11 Herbaspirillum chlorophenolicum AB094401(99%)

12 Rhizobium etli AM921623 (99%); Rhizobiumleguminosarum DQ099745 (98%); Rhizobiumindigoferae AY034027 (98%)

13 Rhizobium leguminosarum EU256425 (100%);14 Sulfolanivorax yambarensis AB285481 (98%)15 Rhizobium sullae EU256432 (99%); Rhizobium

etli EF506199 (99%); Rhizobium gallicumAY509211 (99%)

16 Sinorhizobium meliloti DQ423246 (99%);Sinorhizobium kummerowiae AF364068 (99%)

17 Sinorhizobium meliloti EU271786 (99%);Sinorhizobium kummerowiae AF364068 (99%)

18 Aminobacter aminovorans AF329835 (99%);methylotrophic proteobacterium AF250404(99%)

19 Rhizobium sp. AY141985 (100%);Agrobacterium tumefaciens EU118772 (100%)

20 Paenibacillus sp. EU332823 (98%); Paenibacillusborealis AJ011323 (97%); Paenibacillus odoriferEF199999 (97%)

21 Pseudomonas sp AY835583 (100%)22 Mycobacterium goodii DQ447773 (99%);

Mycobacterium peregrinum AF058712 (98%)23 Bradyrhizobium elkanii DQ485704 (99%);24 Caulobacter sp. AJ227777 (100%); Caulobacter

vibrioides AJ227755 (99%); Caulobactercrescentus AE005673 (99%)

26 Asticcacaulis taihuensis AY500141 (99%);Asticcacaulis benevestidus AM087199 (98%);Asticcacaulis biprosthecium AB014055 (98%)

27 Acidovorax sp AF235010 (100%); Acidovoraxdefluvii Y18616 (98%)

28 Paenibacillus sp AM162331 (99%);29 Lysinibacillus fusiformis EU187493 (100%);

Bacillus sphaericus DQ923481 (100%)30 Paenibacillus polymyxa AY359623 (99%)31 Paenibacillus sp AM162331 (99%);32 Mesorhizobium huakuii EU399717 (99%);

Mesorhizobium amorphae EU399707 (99%);Mesorhizobium plurifarium EU256435 (99%)

Isolate profile numbers in italics indicate non-nodulatingbacteria. Note that a recent molecular systematic study indi-cates that Sinorhizobium and Ensifer should be united into asingle genus (Martens et al. 2007).



2001a,b). It has been argued that reversing thesetrends and managing salinization through improvedsoil structure and lowering of the water table, willrequire the revegetation of many thousands of hectareswith deep-rooted perennial plants (Barrett-Lennard2002; Clarke et al. 2002; Cocks 2003). The spatialscale of replanting required is such that direct drillingis the only cost-effective method of tackling thisproblem and it is imperative that strategies be devel-oped that will improve the success of these efforts.

Integrating the use of salt-tolerant rhizobial isolateswith the most appropriate plant species potentiallyrepresents an important tool for improving the successof revegetation in saline soils. This is particularly thecase, given that rhizobial symbioses promote rapidearly growth and increased survival of native legumesthat may extend beyond host plants to other species,and can significantly impact on long-term viability(Thrall et al. 2005). Salt tolerance has been observedin rhizobia isolated from a range of legume species(Singleton et al. 1982; Elsheik 1998; Mohamed et al.2000; Shamseldin &Werner 2005). Salt-tolerant rhizo-bia have been shown to be more effective than saltsensitive isolates in promoting plant growth of theAfrican species Acacia nilotica under saline conditions(Lal & Khanna 1994); similar results have beenreported for other host species (Zou et al. 1995;Hashem et al. 1998; Shamseldin & Werner 2005). Incontrast, some studies have found no added benefit ofsalt-tolerant rhizobia for plant performance in salinesoils (Lal & Khanna 1994). Inconsistent plantresponses in such studies may be partly due to aninteraction where plant growth responses along salin-ity gradients depend on the degree of salt-tolerance inboth host and symbiont.We have recently shown that,while salt-tolerant specias of Acacia are less responsive

to inoculation overall (Thrall et al. 2008), their growthperformance appears to be greatest when associatedwith more salt-tolerant rhizobial strains (P. H.Thrall &J. Bever 2008, unpubl. data). This suggests that, whileisolates able to survive and grow under highly salineconditions may be prime candidates for inoculantsthat can be used to ameliorate landscape degradationthrough large-scale revegetation programmes, moreresearch is needed to disentangle plant–symbiont rela-tionships along environmental stress gradients.

Preliminary results from glasshouse trials indicate,not only that at least some salt-tolerant strains are alsoeffective at N2-fixation and promotion of host plantgrowth, but that there may be significant ecologicaltrade-offs between these traits (Thrall et al. 2008).Overall, the distribution and diversity of rhizobialspecies depends not only on host species distribution,but also on the diverse environments in which bothhost plant and symbionts have evolved (Parker et al.2002). Thus, the characterization of novel bacterialgenomic species (many unique to saline sites) showsthat the diversity of rhizobia associated with AustralianAcacia spp. is significantly greater than previouslydescribed (Lafay & Burdon 1998, 2001). Importantly,with regard to the potential utility of these rhizobia forlarge-scale revegetation programmes, this study hasrevealed significant differences in growth physiologyand salt tolerance, as well as generic shifts in commu-nity composition relation to soil salinity.

ACKNOWLEDGEMENTS

This work was supported by the National Action Planfor Salinity and Water Quality (project #202749). Wethank D. Millsom, D. Gailey, L. Hyde, J. Spence and

Fig. 5. Two-dimensional principal co-ordinate diagram of PCR-RFLP banding patterns of rhizobial isolates. Capital lettersindicate previously characterized isolates (Lafay & Burdon 1998, 2001), and numbers indicate new profiles identified in thisstudy. White symbols indicate nodulating bacteria, and black symbols indicate non-nodulating taxa.



M. Driver for assistance with collecting soils fromsaline sites, L. Bulkeley, J. McKinnon and L. Li fortechnical support, J. Slattery for help with generatingrhizobial cultures and J. Brockwell for advice with thelaboratory growth experiments.

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