INVERTEBRATE MICROBIOLOGY

Bacterial Community and Nitrogen Fixationin the Red Turpentine Beetle, Dendroctonus valens LeConte(Coleoptera: Curculionidae: Scolytinae)

Jesús Morales-Jiménez & Gerardo Zúñiga &

Lourdes Villa-Tanaca & César Hernández-Rodríguez

Received: 23 April 2009 /Accepted: 27 May 2009 /Published online: 20 June 2009# Springer Science + Business Media, LLC 2009

Abstract The red turpentine beetle, Dendroctonus valensLeConte (Coleoptera: Curculionidae: Scolytinae), colonizesall pines species within its native range throughout Northand Central America. Recently, this species was acciden-tally introduced to China, where it has caused severedamage in pine forests. It belongs to a group of beetles thatspend most of their lives between the tree bark andsapwood, where it feeds on phloem: a poor substrate withvery low nutritional value of nitrogen and toxic propertiesdue to its high content of secondary defensive compounds.The aim of this study was to characterize the bacterialcommunity of the D. valens gut by culture-dependent and -independent methods. Polymerase chain reaction denatur-ing gradient gel electrophoresis and ribosomal gene libraryanalyses revealed that species diversity in the D. valens gutwas relatively low, containing between six and 17 bacterialspecies. The bacterial community associated with larvaeand adults was dominated by members of the followinggenera: Lactococcus, Acinetobacter, Pantoea, Rahnella,Stenothrophomonas, Erwinia, Enterobacter, Serratia, Jan-ibacter, Leifsonia, Cellulomonas, and Cellulosimicrobium.

The members of the last four genera showed cellulolyticactivity in vitro and could be involved in cellulosebreakdown in the insect gut. Finally, nitrogen fixation wasdemonstrated in live larvae and adults; however, capacity ofnitrogen fixing in vitro was not found among enterobacte-rial species isolated in nitrogen-free media; neither werenifD nor nifH genes detected. In contrast, nifD gen wasdetected in metagenomic DNA from insect guts. Theidentification of bacterial species and their potentialphysiological capacities will allow exploring the role ofgut symbiotic bacteria in the adaptation and survival of D.valens in a harsh chemical habitat poor in nitrogen sources.

Introduction

Insects are one of the most successful and diverse groupson earth; they are adapted to a wide variety of diets and livein practically any habitat. Bark beetles have different lifestyles ranging from highly aggressive (tree killing),facultative (colonizing weak or recently killed trees),parasitic (using living trees), to saprophagous (using deadhosts) [50]. In general, they tunnel through the bark toconstruct galleries in the phloem and cambial layers uponwhich they feed [74].

The phloem is a substrate rich in cellulose (22.1% of dryand ethanol-benzene-insoluble material), hemicellulose(15% of dry and ethanol-benzene-insoluble material), andsoluble carbohydrates (120–200 mg g−1) but poor inassimilable nitrogen (11–13 mg g−1 of dry weight) [66,70, 72]. This situation requires bark beetle species to have ahigh phloem consumption rate and to establish symbioticrelationships with microorganisms that supply assimilablecarbon and nitrogen sources and growth factors [3, 5]. Themicroorganisms associated with the insect gut could be

Microb Ecol (2009) 58:879–891DOI 10.1007/s00248-009-9548-2

J. Morales-Jiménez : L. Villa-Tanaca :C. Hernández-Rodríguez (*)Departamento de Microbiología,Escuela Nacional de Ciencias Biológicas,Instituto Politécnico Nacional,Prol. De Carpio y Plan de Ayala. Col. Sto. Tomas,Mexico, Distrito Federal CP 11340, Mexicoe-mail: chdez38@hotmail.com

G. ZúñigaDepartamento de Zoología,Escuela Nacional de Ciencias Biológicas,Instituto Politécnico Nacional,Prol. De Carpio y Plan de Ayala. Col. Sto. Tomas,Mexico, Distrito Federal CP 11340, Mexico

particularly important because their function in the gut mayhelp in food digestion and supply nutrients and growthfactors [63]. Bacterial communities associated with the gutsof bark beetles have not been well investigated, while thegut microbiology of other insect groups has been studiedwidely [8, 45, 48]. Thus, pathogenic [44], cellulolytic [23],nitrogen-fixing [9], and pheromone-producing [7] bacteriahave been isolated of these beetles. In particular, there areonly two works that describe the gut bacterial communityassociated with two bark beetles of the genus Dendrocto-nus: Dendroctonus frontalis (Zimmermann) and Dendroc-tonus micans (Kugelann) [69, 77] using culture-dependentand -independent strategies. These descriptive studiessuggest that bacteria and bark beetles have a mutualismrelationship; however, this relation has not yet beendemonstrated properly.

The red turpentine beetle, Dendroctonus valens LeConte(Coleoptera: Curculionidae: Scolytinae), is found naturallythroughout North and Central America, except in the SEUSA [32]. This species is parasitic—it lives primarily onliving trees without causing mortality. It is occasionally asecondary pest, killing trees infested by other bark beetlesor damaged by fire [18, 74]. Recently, this species wasimported from North America to China [19], where it iskilling trees of normal or near-normal vigor of Pinustabuliformis [41, 76]. Despite the wide geographic distri-bution, wide host range, and the recent economical andecological importance of D. valens, little is known aboutthe potential participation of the bacterial gut community innitrogen fixation, cellulose digestion, or other roles in hostbeetle nutrition. The aim of this study was to characterizeand to identify the bacterial community of D. valens gut byculture-dependent methods and culture-independent molec-ular techniques. In addition, we wished to documentpotential capacities of the bacteria related to the cellulolyticactivity and nitrogen fixation which may be important inthe nutritional ecology of bark beetles.

Materials and Methods

Beetle Collection

D. valens adults were collected from four geographicallocations in Mexico (Table 1). Insects were obtaineddirectly from galleries of infested pine trees using fineforceps and during the first stages of colonization. Inaddition, ultimate instar larvae were collected forcomparative purposes. Live adults and larvae weretransported to the laboratory in sterile vials containingmoist paper. The insects were disinfected superficiallywith 70% ethanol and submerged repeatedly in aphosphate buffer solution (PBS) to avoid external

contamination. Insects were dissected under sterileconditions, and the gut extraction was performed bythe elimination of elytra, wings, and tergites to leave theinsect abdomen exposed. The guts were transferredindividually to 1.5-ml microcentrifuge tubes with0.2 ml of PBS or culture medium.

Isolation of Bacteria

Individual guts from larvae and adults were placedseparately in sterile microcentrifuge tubes, where they werecrushed in 200 µl of PBS. Serial tenfold dilutions werespread on duplicate plates of Luria–Bertani agar and asimple medium, previously described by Bridges [9]. Plateswere incubated at 28 °C for 2 to 5 days; each colony typewas categorized and counted. Pure cultures were obtainedand stored at −70 °C for further analysis.

Differentiation of Bacteria by Randomly AmplifiedPolymorphic DNA Fingerprints

To extract DNA, pure colonies were transferred to tubeswith 5 ml of Luria–Bertani medium, incubated at 28 °C for2 days [22]. Based on randomly amplified polymorphicDNA (RAPD)-polymerase chain reaction (PCR) finger-prints generated with one primer, different bacterial groupswere identified [73]. The reaction mixture contained0.2 µM of dNTPs, 2.5 µl 1× PCR buffer, 3 mM MgCl2,0.8 pM primer, 1.5 U of Taq polymerase (Invitrogen LifeTechnologies, Sao Paulo, Brazil), and 10–100 ng ofbacterial DNA adjusting to 25 µl with sterilized deionizedwater. The reaction conditions were 94 °C for 5 min; 35cycles of 60 s at 94 °C, 60 s at 38 °C, and 60 s at 72 °C;and a final extension at 72 °C for 10 min. At least twoisolates of each group were selected for 16S rRNAsequencing.

PCR of 16S rRNA Gene Fragments

This gene was amplified by PCR using primers 8 forward( 5 ′ - GCGGATCCGCGGCCGCTGCAGAGTTTGATCCTGGCTCAG-3 ′) and 1,492 reverse (5 ′-GGCTCGAGCGGCCGCCCGGGTTACCTTGTTACGACTT-3′) [55]. Final concentrations for 25 µl PCRreactions were as follows: 10 ng of metagenomic DNAisolated from gut, 0.8 pM of each primer, 0.2 mM dNTPs,2.5 mM MgCl2, 1 U of Taq polymerase, and 1× Taqpolymerase buffer (Invitrogen Life Technologies, SaoPaulo, Brazil). The reaction conditions were 94 °C for7 min; 35 cycles of 60 s at 94 °C, 60 s at 55 °C, and 60 s at72 °C; and a final extension at 72 °C for 7 min. PCRproducts were purified using the QIAquick PCR purifica-tion kit (Qiagen, Valencia, CA, USA) and sequenced in an

880 J. Morales-Jiménez et al.

ABI PRISM 310 genetic analyzer (Applied Biosystems,Foster City, CA, USA) using the same primers.

16S rRNA Gene Libraries

Complete guts from five larvae or adult beetles were pooledand placed in 1.5 ml microcentrifuge tubes containing200 µl PBS and stored at −20 °C until metagenomic DNAextraction was done. Guts were macerated with a plasticpestle and vortexed at medium speed for 10 s. The DNAextraction and 16S rRNA amplification conditions weresimilar to those described above. PCR products werepurified using the QIAquick PCR purification kit (Qiagen)and cloned in Escherichia coli Top-10 cells (Invitrogen LifeTechnologies, Carlsbad, CA, USA) with the PCR® 2.1-Topo® vector (Invitrogen-Life Technologies, Carlsbad, CA,USA) according to manufacturer's instructions. Transform-ants were subjected to plasmid extraction by standardmethods [57], and a restriction analysis with EcoRI wasperformed to detect insertions. Plasmidic DNA of eachclone of the 16S rRNA library was digested withendonuclease HpaII to display restriction fragment lengthpolymorphism (RFLP) patterns by electrophoresis in 3%high-performance agarose 1000 (GIBCO Laboratories,Grand Island, NY, USA). The plasmids of each RFLPpattern were extracted using High Pure Plasmid IsolationKit (Roche) and sequenced with the ABI PRISM 310genetic analyzer (Applied Biosystems) using M13 primers.

Phylogenetic Analyses

Clone sequences obtained were tested for chimera struc-tures by using RDP Check Chimera (http://35.8.164.52/cgis/chimera.cgi?su=SSU) [20], and chimeras were exclud-ed from further analysis. Sequences from clones andisolated bacteria were compared with the nonredundantGenBank library using BLAST search [2]. A collection oftaxonomically related sequences were obtained from theNational Center for Biotechnology Information Taxonomy

Homepage (http://www.ncbi.nlm.nih.gov/Taxonomy/taxo-nomyhome.html/) and Ribosomal Database Project-II Re-lease 9 (http://rdp.cme.msu.edu). DNA sequences werealigned using CLUSTAL X [65] and edited and confirmedvisually in BIOEDIT [34].

Maximum likelihood analyses were performed usingPhyML [33; http://atgc.lirmm.fr/phyml/]. MODELTEST3.06 [51] was used to select appropriate models of sequenceevolution by the AIC model [52]. The GTR + I +G model(α=0.442 for the gamma distribution; A=0.23, C=0.23,G=0.31, T=0.22; p-inv=0.26) was selected for the treesearch. The confidence at each node was assessed by 1,000bootstrap replicates [36]. Anabaena affinis was used asoutgroup. The similitude percentages among sequenceswere calculated using MatGAT v. 2.01 software [14]. Thelimits for genus and species were set at 95% and 97%,respectively [60]. Sequences of this study were deposited inthe GenBank database under the accession numbers:FJ811851 through FJ811890 (Table 2).

To assess the richness in a community, Chao1 richnessestimator was calculated using DOTUR [60]. This nonpara-metric richness index estimates the diversity of a communitybased on the number of singletons or operational taxonom-ical units (OTUs) represented by only one sequence anddoubletons (OTUs represented by two sequences) found in asample [6]. Terminal Chao1 richness estimates are reportedfor 3% difference between sequences.

Denaturing Gradient Gel Electrophoresis Analysis

To know whether bacterial communities change across gutregions, denaturing gradient gel electrophoresis (DGGE)analysis was performed. Five adult guts were separated intoanterior and posterior midgut and hindgut. Each region wasstored frozen at −20 °C until DNA extraction was done asdescribed above. Primers P3 (5′-CGCCCGCCGCGCGCGGCGGGCGGGGCGGGGGCACGGGGGGCCTACGGGGGCAGCAG-3′) and P2 (5′-ATTACCGCGGCTGCTGG-3′) were used for the 3-V region of 16S rRNA

Table 1 Red turpentine beetle (D. valens) samples used in this study

Location Locationcode

Latitude/longitude

Host tree No. ofinsects

San Juanito, Bocoyna, Chihuahua BCH 27°55′ N Pinus arizonica 20 adults107°55′ W

El Wicle, Nuevo Casas Grandes, Chihuahua WCH 27°55′ N Pinus engelmannii 45 adults107°55′ W

Centro Interdisciplinario de Ciencias de la salud (CICS-IPN),Milpa Alta, D.F.

CDF 19°04′ N Pinus montezumae 35 larvae98°58′ W

Los Pozos, Gómez Farias, Jalisco. GFJ 19°88′ Pinus montezumae or Pinusleiophylla

25 adultsN 103°40′ W

Bacterial Community and N Fixation in the Red Turpentine Beetle 881

amplification [46]. The reaction mixture contained0.2 µM of dNTPs, 2.5 µl 1× PCR buffer, 3 mM MgCl2,0.8 pM primer, 1.5 U DNA Taq polymerase (InvitrogenLife Technologies, Sao Paulo, Brazil), and 10 ng of DNAadjusting to 25 µl with sterilized deionized water. The

reaction conditions were 94 °C for 5 min; 35 cycles of 60 sat 94 °C, 60 s at 55 °C, and 60 s at 72 °C; and a finalextension at 72 °C for 10 min. The expected size of theamplified fragment was 240 bp. DGGE analysis wasperformed with a DCode mutation detection system (Bio-

Table 2 GenBank accession numbers assigned to the sequences generated in this study

Isolated strain or ribosomal library clone code (GenBank accessionnumber)

Closest species in GenBank (GenBank accessionnumber)

Similarity(%)a

3 BCH (FJ811856) Rahnella aquatilis (U970757) 98

6 BCH (FJ811855) 97.9

7 BCH (FJ811854) 97.9

9 BCH (FJ811853) 97.7

13 BCH (FJ811852) 97.9

17 BCH (FJ811857) 98

2 WCH (FJ811858) 98.1

3 WCH (FJ811860) 97.7

2B CDF (FJ811859) 98.1

C7 WCH (FJ811883) 97.3

C15 WCH (FJ811882) 97.4

C4 WCH (FJ811884) 97

C2 WCH (FJ811885) 96.9

2A CDF (FJ811861) Serratia proteamaculans (AY040208) 99.8

3 CDF (FJ811862) 99.7

2 BCH (FJ811864) Serratia liquefaciens (AJ306725) 99.7

15 BCH (FJ811863) 99.7

1 BCH (FJ811865) 99.5

19 CDF (FJ811866) 99.2

5 WCH (FJ811874) Enterobacter aerogenes (AJ251468) 98.6

1 WCH (FJ811873) 98.8

11 WCH (FJ811872) 98.8

7 WCH (FJ811871) 98.8

1 CDF (FJ811870) Pantoea dispersa (AB273743) 96.5

MP1 GFJ (FJ811869) Erwinia mallotivora (AJ233414) 96.7

MP1FX GFJ (FJ811868) 96.8

16 CDF (FJ811867) Pantoea cedenensis (AF130971) 99.4

CP20 WCH (FJ811886) Pantoea dispersa (AB273743) 96.9

8 CDF (FJ811876) Acinetobacter lwoffii (DQ328322) 98.5

MA3B GFJ (FJ811875) Acinetobacter haemolyticus (AM184255) 97.5

CP23 WCH (FJ811887) Acinetobacter schindleri (FJ373024) 98.8

4 CDF (FJ811851) Stenotrophomonas maltophilia (EF620448) 99.7

MA2 GFJ (FJ811877) Leifsonia shinshuensis (DQ232614) 98.8

PL2 GFJ (FJ811881) Cellulomonas xylanilytica (AY303668) 98.7

MA3A GFJ (FJ811880) Cellulosimicrobium cellulans (AY665978) 98.2

PL5 GFJ (FJ811879) 99.2

MA1B GFJ (FJ811878) Janibacter melonis (AY522569) 99.6

CP4 WCH (FJ811884) Lactococcus lactis (EU104368) 99.3

CP1 WCH (FJ811888) 99.3

C16 WCH (FJ811890) 99.3

a The similitude percentages among sequences were calculated using MatGAT v. 2.01 software [14].

882 J. Morales-Jiménez et al.

Rad Laboratories, Hercules, CA, USA). Gels of 8%acrylamide (37:1 acrylamide–bisacrylamide) were formedwith a linear gradient between 20% and 80% denaturant;100% denaturant is defined as 7 M urea and 40% (v/v)formamide. Gels were run at 60 V for 16 h and maintainedat 60 °C during 16 h in 1× TAE buffer (40 mM Tris,20 mM acetate, 1 mM EDTA). DNA bands were visualizedby a previously described silver stain procedure [58].

Isolation of Nitrogen-Fixing Bacteria

Isolation of potential nitrogen-fixing bacteria was carriedout at 28 °C during at least 72 h in a nitrogen-free mediumdescribed by Bridges [9]. The content of the gut from liveadults and larvae was inoculated separately in liquidmedium. Three successive subcultures into fresh mediumwere performed to be sure that the isolated bacteria cangrow in this nitrogen-free medium. Finally, bacteria wereisolated in nitrogen-free solid medium and cultured in ananaerobic container system (GasPak™ EZ, BD, Maryland).

Acetylene Reduction Assay

Acetylene reduction, as a functional assay for nitrogenfixation, was performed in live adults and larvae of D.valens immediately after collecting them. Individual larvaeor adults were incubated in closed vessels (10 ml) contain-ing a wet sheet of paper with an acetylene-enrichedatmosphere. Acetylene was produced by dissolving calciumcarbide in tap water and injected to a final concentration of1% (v/v) by replacement of an identical volume of air.Autoclaved insects were used as negative control. Nitroge-nase activity was determined by measuring the acetylenereduction activity of individual larvae or adults in closedvessels with a Varian 3300 (Walnut Creek, CA, USA) gaschromatograph with a flame ionization detector. Acetylenereduction activities with isolated bacteria were performedusing WATC-4, Winodrasky, and Bridges semisolid media[4, 9, 35] inoculated with a single colony of each bacterialstrain. The media were incubated for 24 h at 28 °C, andthen acetylene-enriched atmospheres to a final concentra-tion of 1% and 10% were injected. The acetylene reductionactivity was measured as above.

Isolation of Cellulolytic Microorganisms

Individual guts from adults and larvae were placed in sterileEppendorf tubes containing 200 µl of PBS and processed asdescribed in previous sections. Serial tenfold dilutions werespread on duplicate plates of Congo red agar (0.5 g l−1

K2HPO4, 0.25 g l−1 MgSO4, 1.88 g l−1 powder cellulose,0.2 g l−1 Congo red, 2 g l−1 gelatin, 100 ml l−1 soil extract,15 g l−1 agar). Plates were incubated in a growth chamber at

28 °C for 3 to 5 days. The cellulolytic activity ofmicroorganisms was detected by a clear zone around thecolonies [64]. Pure cultures were obtained by multiplesubsequent subculturing on of Congo red agar. Enzymeactivity was indexed as the diameter of the colony plus theclear zone divided by the diameter of the colony. Twodiameters of at least two colonies from the same bacterialisolate were measured [23]. To confirm the cellulolyticactivity of these isolates, we determined reducing sugarsreleased from the cellulose powder after 24 h of growth at28 °C in Congo red free Cellulose media. Cultures werecentrifuged at 10,000×g for 10 min at 4 °C, and thesupernatant was stored (1 to 4 weeks) at −20 °C untilenzyme activities were measured. The supernatant (0.5 ml)was incubated in 0.5 ml powdered cellulose (0.5%) in0.05 M sodium phosphate buffer. All enzyme reactionswere carried out in duplicate at 37 °C for 60 min. Reducingsugars were measured by the 3,5-dinitrosalicylic acidmethod [42] using glucose as standard. Units of activityare defined as micromoles of glucose released per minute(IU).

Results

Culture-Dependent and -Independent Analysis of BacterialCommunity

A total of 2.11×106 CFU/gut of heterotrophic culturablebacteria in nutritive agar was estimated in the whole gut ofadult insects. D. valens harbored 0.41±0.0621, 0.34±0.056, and 1.35±0.067×106 CFU in the anterior midgut,posterior midgut, and hindgut, respectively. The hindgutmaintained the largest bacterial populations with 63% oftotal culturable bacteria. From 98 bacterial strains, 24RAPD profiles were recognized. The phylogenetic analy-sis of 16S rRNA of the RAPD profiles identified 11bacterial genera including 13 bacterial species (Table 3and Fig. 1).

On the other hand, the evaluation of gut bacteria byculture-independent analysis revealed a low bacterialdiversity, since only 12 HpaII RFLP patterns of 16S rRNAlibrary were identified belonging to four bacterial speciesaccording to the phylogenetic analysis (Table 3 and Fig. 1).These species were affiliated with known taxa of γ-Proteobacteria and low G+C Gram-positive bacterialdivisions. Adult beetle guts harbored Rahnella aquatilis,Lactococcus lactis, Acinetobacter schindleri, and Pantoeadispersa with relative abundances of 23.07%, 38.47%,19.23%, and 19.23%, respectively.

The rarefaction analyses of 16S rRNA libraryconfirmed the presence of a relatively scarce bacterialcommunity in adult beetle guts. Rarefaction curve had a

Bacterial Community and N Fixation in the Red Turpentine Beetle 883

low slope with a 3% level of difference amongsequences, indicating that most of the species had beensampled. At 10% and 20% level of differences amongsequences, the curves were mostly flattened, suggestingthat most of the classes and phyla associated with adultbeetle gut were represented. The terminal Chao1richness estimates for the library was five OTUs, where

an OTU was defined as a group of sequences differingby no more than 3%.

DGGE analysis of bacterial community showed a totalof 12 bands in adult females and eight in males. There wereno major differences between anterior midgut and hindgut;however, the hindgut exhibited more bands than themidgut. The comparison of migration bands of gut

Table 3 Bacterial taxa associated with guts of larvae and adults of the red turpentine beetle in culture-dependent and culture-independent analyses

Different isolate RAPD or cloneRFLP patterns

Phylogenetic relationships Microbialgroupaffiliationa

Insect stage

Bacterialdivision

Genus (≥95%identity)

Database matches(≥97% identity)

Larvae Adults

Identified by culture dependent analysisb

3 BCH, 6 BCH, 7 BCH, 9 BCH, 13BCH, 17 BCH, 2 WCH, 3WCH, 2B CDF

γ-Proteobacteria

Rahnella R. aquatilis (U970757) R. aquatilis + +

2A CDF, 3 CDF Serratia S. proteamaculans(AY040208)

S.proteamaculans

+

2 BCH, 15 BCH, 1 BCH, 19 CDF Serratia S. liquefaciens(AJ306725)

S. liquefaciens + +

5 WCH, 1 WCH, 11 WCH, 7 WCH Enterobacter Uncultured bacteriumclone 6 s2(DQ068850)

E. aerogenes +

1 CDF Pantoea Pantoea sp. +

MP1 GFJ, MP1FX GFJ Erwinia Uncultured bacteriumclone spb28c6(DQ321565)

Erwinia sp. +

16 CDF Pantoea P. cedenensis(AF130971)

P. cedenensis +

8 CDF Acinetobacter A. lwoffii (DQ328322) A. lwoffii +

MA3B GFJ Acinetobacter Uncultured bacteriumclone (AF534208)

A. haemolyticus +

4 CDF Stenotrophomonas S. maltophilia(EF620448)

S. maltophilia +

MA2 GFJ Actinobacteria Leifsonia L. shinshuensis(DQ232614)

L. shinshuensis +

PL2 GFJ Cellulomonas C. xylanilytica(AY303668)

C. xylanilytica +

MA3A GFJ, PL5 GFJ Cellusimicrobium C. cellulans(AY665978)

C. cellulans +

MA1B GFJ Janibacter J. melonis (AY522569) J. melonis +

Identified by culture independent analysisc

C7 WCH, C15 WCH, C4 WCH γ-Proteobacteria

Rahnella Uncultured bacteriumclone spb28a3(DQ321557)

R. aquatilis +

C2 WCH Uncultured bacteriumclone spb28a3(DQ321557)

Rahnella sp. +

CP20 WCH Pantoea Pantoea sp. +

CP23 WCH Acinetobacter A. schindleri(FJ373024)

A. schindleri +

CP4 WCH , CP1 WCH, C16 WCH Firmicutes Lactococcus L. lactis (EU104368) L. lactis +

BCH San Juanito, Bocoyna, Chihuahua; WCH El Wicle, Nuevo Casas Grandes, Chihuahua; CDF Centro Interdisciplinario de Ciencias de la salud(CICS-IPN), Milpa Alta, D.F.; GFJ Los Pozos, Gómez Farias, Jaliscoa The limits for genus and species were 95% and 97%, respectively [60]b Each one represents a different RAPD pattern generated using the prime OPB-01c Each one represents a different RFLP pattern defined with endonuclease HpaII

884 J. Morales-Jiménez et al.

7 BCH9 BCH13 BCH6 BCH3 BCH17 BCH2 WCH

Rahnella aquatilis (U90757)Uncultured bacterium clone spb28a3 (DQ321557)

2B CDF3 WCH

C7 WCHC15 WCHC4 WCHC2 WCH

Serratia plymuthica (AJ233433)Serratia grimesii (AF286868)2A CDF3 CDFSerratia proteamaculans (AY040208)

Serratia liquefaciens (AJ306725)2 BCH15 BCH1 BCH19 CDFUncultured bacterium clone rRNA198 (AY958971)

Enterobacter intermedius (AF310217)Enterobacter aerogenes (AJ251468)Uncultured bacterium clone 6s2 (DQ068850)5 WCH1 WCH11 WCH7 WCH

Pantoea dispersa (AB273743)1 CDFCP20 WCH

Erwinia mallotivora (AJ233414)Uncultured bacterium clone spb28c6 (DQ321565)

MP1 GFJMP1FX GFJ

Erwinia toletana (AF130968)16 CDFPantoea cedenensis (AF130971)

Pseudacinetobacter hongkongensis (AF543466)Acinetobacter lwoffii (DQ328322)

8 CDFAcinetobacter rhizosphaerae (DQ536511)

Acinetobacter brisoui (DQ832256)Vestimentiferan symbiont (AB042404)

Uncultured bacterium clone (AF534208)MA3B GFJ

Acinetobacter haemolyticus (AM184255)Acinetobacter schindleri (FJ373024)CP23 WCH

Acinetobacter johnsonii (EF114343)Cibimonas vasta (DQ846687)4 CDFStenotrophomonas maltophilia (EF620448)

Stenotrophomonas rhizophilia (AJ293463)Stenotrophomonas dokdonensis (DQ178977)

Xanthomonas campestris (AY605124)Stenotrophomonas maltophilia (FJ418173)

Leifsonia poae (AM410682)Leifsonia xyli (AJ717351)Leifsonia shinshuensis (DQ232614)MA2 GFJ

Clavibacter michiganensis (D45059)Cellulmonas humilata (X82598)

Cellulomonas humilata (X82449)Cellulomonas xylanilytica (AY303668)PL2 GFJMA3A GFJ

PL5 GFJCellulosimicrobium cellulans (AY665978)

Ornithinicoccus hortensis (AB098587)Janibacter terrae (AF176948)Janibacter anophelis (AY837752)

Janibacter marinus (AY533561)Janibacter melonis (AY522569)MA1B GFJ

Lactococcus piscium (DQ343754)Lactococcus garvieae (AB267897)CP4 WCHCP1 WCHC16 WCHLactococcus lactis (EU104368)

Anabaena affinis (AF247591)

670

574

752

584

783

593

931

503

573

985

897

999

950801

620

981

10001000

750

823

998923

824

837

641

1000

1000

675

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992627

995576

1000

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999

6431000

1000

1000

555938

979

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659

7501000

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717

640913

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517

673996

690

1000

0.05

γ-P

rote

obac

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aA

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obac

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Figure 1 Maximum likelihood tree (-lnL = 15344) of bacterialcommunity associated with D. valens gut. The 16S rRNA sequence ofA. affinis was used as outgroup. Scale bar indicates 5% estimated

sequence divergence. Bootstrap support values are indicated for majornodes having values of ≥50%

Bacterial Community and N Fixation in the Red Turpentine Beetle 885

metagenomic DNA and isolated bacteria by DGGE analysisrevealed that R. aquatilis and Enterobacter aerogenes werelargely dominant species (Fig. 2b). The estimation of theband intensity and DGGE patterns from different gutregions suggest that both species are broadly distributedin the alimentary canal from males and females (Fig. 2a).

Some γ-Proteobacteria and Actinobacteria, species thatwere cultured were not detected in culture-independentanalyses; however, Acinetobacter, Rahnella, and Pantoeawere found in D. valens gut using both methods. On theother hand, Lactoccocus was found exclusively in culture-independent analyses. The evaluation of the midgut andhindgut bacteria of D. valens, using both culture-dependentand -independent methods, allowed identification of 17bacterial species from γ-Proteobacteria, Firmicutes, andActinobacteria groups.

Acetylene Reduction in the Red Turpentine Beetle

A significant acetylene reduction to ethylene wasdetected in live larvae and adult insects (15.96±2.22;12.59±3.39 nmol ethylene/h/insect, respectively), but noactivity was present in dead larvae or adult insects or ina negative control (1.12±0.47; 0.34±0.22; 0±0 nmolethylene/h/insect, respectively; one-way analysis ofvariance: F=15.09; P<0.05; Fig. 3). The nitrogen fixedper live larvae or adults per day was equivalent to 5.6 and

4.4 µg, respectively. Moreover, bacterial nifD genefragment was amplified from gut regions. These datasuggest that nitrogen fixation occurs in D. valens gut. R.aquatilis, Stenotrophomonas maltophilia, Pantoea cede-nensis, and P. dispersa were isolated in nitrogen-freemedium. Although a visible growth of these bacteria wasclearly observed, no in vitro acetylene reduction wasdetected in different nitrogen-containing and -free mediausing several carbon sources.

nmol

eth

ylen

e pe

r in

sect

per

hou

r

0

2

4

6

8

10

12

14

16

18

20

L.L . L.A. D.L. D.A. C.

Figure 3 Nitrogenase activity as estimated by acetylene reduction oflive D. valens larvae and adults. Negative controls: 10% acetylenewithout beetles and 20% acetylene with autoclave-killed beetles (L.L.live larvae, L.A. live adults, D.L. dead larvae, D.A. dead adults, Ccontrol without insects)

1 2 3 4 5 6 1 2 3 4 5 6 7 8 9 10 11 12

a b

Figure 2 Denaturing gradient gel electrophoresis of 16S rDNA PCRproducts obtained from a field-collected adult guts of D. valens. Adultfemale guts (lane 1 anterior mesenteron, lane 2 posterior mesenteron,lane 3 hindgut) and adult male guts (lane 4 anterior mesenteron, lane5 posterior mesenteron, lane 6 hindgut); b field-collected adult guts ofDendroctonus vales and some bacteria isolated from adult guts on

different culture media: adult female guts (lane 1 anterior mesenteron,lane 2 posterior mesenteron, lane 3 hindgut), adult male guts (lane 4anterior mesenteron, lane 5 posterior mesenteron, lane 6 hindgut),bacterial isolates (7, 10–12 E. aerogenes; 8, 9 R. aquatilis). Openblack arrow common bands in all gut regions and sexes. Full blackarrow R. aquatilis. Black arrowhead E. aerogenes

886 J. Morales-Jiménez et al.

Cellulolytic Bacteria

The density of aerobic cellulolytic bacteria was 900±210 CFU/gut. The number of cellulolytic bacteria waslower than the total number of heterotrophic aerobiccultured bacteria (2.11×106 CFU/gut). The Actinobacteria,Janibacter melonis, Leifsonia shinshuensis, Cellulomonasxylanilytica, and Cellusimicrobium cellulans, were isolatedand identified from D. valens gut (Table 3). All of themshowed cellulose degrading activity (Fig. 4 and Table 4).

Discussion

Bacterial communities associated with insect guts havebeen widely studied [11, 31, 45, 49] and have been found tobe variable and astonishingly diverse. For example, intermites, cockroaches, and scarab beetles, >100 bacterialspecies have been reported from the gut only [30, 31, 37,48]; however, in broad-headed bugs, the number reported ismuch lower like just one genus [40]. The bacterial diversityassociated with D. valens appears to be unusually lowcompared to wood-feeding termites. Particularly, the hind-

gut of termites contains a radial and axial structure withgradients of metabolites and physicochemical conditions[12]. In addition, bark beetles and symbionts experiencetoxic compounds [13], such as monoterpenes, early in theirlife cycles, which could limit the presence of many bacterialspecies. As a consequence, the termite gut bacterialcommunity is more dense, complex, and metabolicallydiverse [71] than bacterial community of D. valens.Apparently, the bacterial diversity harbored in the gut ofinsects depends on the host's lifestyle, habitat, and diet. Thefew studies carried out in bark beetles have shown that gutbacterial diversity is low [69], which may be due to the lownutritional value of phloem, a substrate with a high C/Nratio (43–52) [70] and limited essential amino acids.

The results of this study showed that the D. valens gut islow in bacterial diversity, since only 17 species wereidentified using both16S rRNA and DGGE. These resultsagree with studies on D. frontalis, D micans, and Ips pini,which also found a low diversity using the same strategies[24, 69]. While some bacterial genera, such as Acineto-bacter, Enterobacter, Klebsiella, Pantoea, Rahnella, andStenotrophomonas, are common in the guts of D. valensand D. frontalis, only three species (S. maltophilia, R.

Table 4 Bacterial isolates from D. valens gut with cellulolytic activity

Isolate Alimentary canal region Enzyme activity (indexa/mUI∙ml−1 b)

Cellulomonas xylanilytica (PL2 GFJ) Hindgut 2.7/14.9

Cellulosimicrobium cellulans (PL5 GFJ ) Hindgut 2.9/12.2

Janibacter melonis (MA1B GFJ) Anterior midgut 3/n.d.

Leifsonia shinshuensis (MA2 GFJ) Anterior midgut 3.1/11.5

Cellulomonas xylanilytica (MA3A GFJ) Anterior midgut 2.4/16.4

a Enzyme activity was indexed as the diameter of the colony plus the clear zone around it divided by the diameter of the colony in Congo redcellulose mediumb Each value is the mean of the enzyme activity in two different cultures

Figure 4 Cellulolytic activityof Actinobacteria isolates inCongo red-cellulose. a J. mel-onis MA1B GFJ, b C. xylanily-tica PL2 GFJ, c C. cellulansPL5 GFJ

Bacterial Community and N Fixation in the Red Turpentine Beetle 887

aquatilis, and E. aerogenes) are shared among them.Likewise, bacterial genera Pantoea, Erwinia, and Steno-trophomonas are present in the D. valens and I. pini gut[24]. The presence of Stenotrophomonas and Pantoeagenera in guts of all bark beetles thus far studied suggeststhat members of these genera could play essential andspecific functions in scolytine guts.

Some bacterial groups found in a bark beetle specieshave not been found in others [24, 69]. For example,bacteria such as Rhodobacter sp. and Mycoplasma sp. (α-Proteobacteria) and Leuconostoc mesenteroides, Lactoba-cillus sp., Bacillus cereus, and Enterococcus faecalis(Firmicutes), found in D. frontalis [23, 69] were notidentified in D. valens gut. In contrast, some bacteriaindentified in this study, such as Actinobacteria (J. melonis,L. shinshuensis, C. cellulans, and C. xylanilytica) and L.lactis were not detected in those studies. All bacterialspecies found in D. valens gut have been isolated in guts ofother insects [see for example 11, 15, 29–31, 59, 61, 75].

We did not find obligate anaerobic bacteria in the gut ofD. valens, which agrees with reports in D. frontalis and I.pini [24, 69]. Anaerobic bacteria have been frequentlyfound in the gut of other insects [11, 31, 45, 48]. To ourknowledge, no studies to determine O2, H2, CO2 concen-trations, and redox potential of bark beetle gut have beenperformed, but the absence of anaerobe species may beexplained by the lack of strict anoxic conditions in the barkbeetle gut.

A high abundance of heterotrophic culturable bacteriahas also been observed in other insects, such as termites,crickets, cockroaches [8, 17, 21, 68]. Although the highabundance of bacteria in the hindgut of these groups hasbeen associated to key metabolic processes (e.g., cellulosebreakdown, nitrogen fixation, methanogenesis, acetogene-sis, etc) linked to host nutrition, this may not be the case forthe hindguts of bark beetles. Histological and morpholog-ical studies have shown that the bark beetle hindgut is notan appropriate region where absorption and metabolic

processes take place due to the presence of a nonpermeablechitinous layer in epithelial cells [25–27]. We hypothesizethat the abundance of bacteria in the hindgut of D. valensmay be related to α-pinene transformation sequestered byinsects during their feeding or be involved in the metabo-lism of terpene alcohols, such as trans- and cys-verbenoland myrtenol. This argument is supported by workconducted with Dendroctonus ponderosae [38], whichshowed that symbiotic bacteria can regulate the levels ofthe aggregation pheromones because bacteria may metab-olize α-pinene or their terpene alcohols and produceantiaggregation pheromone, verbenone.

R. aquatilis was the most common species isolated in D.valens (this study) and D. frontalis [23, 69]. Species of thegenus Serratia have been also reported in guts of D.frontalis and D. micans [69, 77] and have been described asinsect pathogens [43, 44]. Also, Pseudomonas spp. werefound in D. frontalis and D. micans, but no species of thisgenus were detected in the gut of D. valens. Pseudomonasspp. of D. micans could have a nutritional role in the gut[77], but to our knowledge, no experimental evidencesupports this proposal. The recurrent isolation of R.aquatilis, E. aerogenes, Pantoea sp., and Serratia spp.from bark beetle guts [10, 43, 44, 69, 77] suggests thatthese bacteria are a constant fraction of the bacterialcommunity maintaining important symbiotic roles withthese insects. For example, facultative anaerobic bacteriafound in this study (Serratia spp., Erwinia spp., and E.aerogenes) could be participating in anaerobic micrositegeneration to allow nitrogen fixation and carbohydratefermentation, as demonstrated for Enterobacter isolatedfrom termites gut, which were implicated in the removal ofoxygen permeated from the exoskeleton, protecting thestrict anoxic conditions of the gut that are essential forcellulose digestion [1].

A taxonomic group especially abundant in ribosomallibraries was L. lactis. These Firmicutes have not beendetected in other bark beetles. Lactic bacteria are abundant in

N2 NH4Nitrogen fixation

Cellulose GlucoseBreakdown

Nitrogen-fixing bacteria

Cellulolytic bacteria

MonoterpenesOxidated

monoterpenes(Semiochemicals)

BiotransformationBacteria

FermentableSugars

CO2, H2

Redox potentialAlcohols and organic acids

production

Facultive anaerobicbacteria

Fermentation

Functions of the bacterial community

Dendroctonus valens

Gut

Figure 5 Putative roles of the bacterial community associated with the gut of D. valens

888 J. Morales-Jiménez et al.

termites, and it has been suggested that they are involved inuric acid recycling [53, 54]; however, a possible function inbark beetles is a highly speculative exercise.

Pantoea species identified in this work have not beenpreviously described in other Dendroctonus species. WhilePantoea agglomerans isolated from D. frontalis andDendroctonus terebrans gut is a nitrogen-fixing bacterium[9, 10, 69], P. dispersa and P. cedenensis isolated in thisstudy were unable to grow in media without a nitrogensource or to reduce acetylene and apparently have no nifDgenes (data not shown). On the other hand, in vegetablemodels, it has been shown that S. maltophilia and Raquatilis participate in nitrogen fixation [4], suggesting thatthese bacterial species found in the gut of D. valens and D.frontalis [69] could have the same role. However, in spitethat both species exhibited a clear growth in liquidnitrogen-free media after at least three successive transfers,no acetylene reduction was detected. Independently fromthis, our results suggest that some nitrogen-fixing activityexists in the D. valens gut, but Ayres et al. [3] indicated thatnitrogen-fixing bacteria in the D. frontalis gut do not satisfycompletely the nitrogen requirements of this species due totheir low abundance and because they have a limitedmetabolic activity. In our opinion, more studies must beperformed to evaluate the proportional contribution ofbacteria to total nitrogen requirements of bark beetlesbecause R. aquatilis was the most abundant bacterium inlarval and adult gut from D. valens. In addition, because R.aquatilis is a conifer endophytic bacterium [16, 39], itmight be easily acquired by bark beetles during theirfeeding. Typing analyses of the strains will be necessary todetermine whether the isolates from conifer phloem belongto the same ecotype as the isolates from insects or whetherthese bacteria are permanent residents of the insect gut.

This is the first report of actinobacterial species (C.cellulans, C. xylanilytica, L. shinshuensis, and J. melonis)isolated from larval and adult gut of D. valens exhibitingcellulolytic properties in vitro. The ability to degradecarboxymethylcellulose of C. cellulans and C. xylanilyticahas been reported previously [56, 62], but this characteristichas not been described in L. shinshuensis and J. melonis.Cellulolytic bacteria were not detected in D. frontalis gut,and apparently, no degradation of cellulose by microbialactivities has been observed [23]. Cellulolytic bacteria inadult D. valens correspond to around 0.04% of the bacterialculturable fraction in Luria–Bertani agar. In other insects,for example in the wood-feeding cockroach (Eublaberusposticus), these bacteria reach 1% of the total culturedbacteria [21]. Cellulose digestion requires a concertedaction of a community of microorganisms with comple-mentary hydrolytic abilities [67]. Possibly, Actinobacteriaconstitute a bacterial community that contributes to thedegradation of cellulose within the insect gut. However, the

presence of these bacteria in the gut of D. valens do notprove the process actually occurs within insect gut.Additional experiments must be performed to confirm thatthe cellulolytic bacteria detected in this work contribute tothe insect diet.

The survival of Dendroctonus in a nitrogen-poorsubstrate requires the intake of supplementary nitrogensources. An important set of evidence supports the ideathat mycangial filamentous fungi contribute to nitrogento the bark beetle diet; however, in non-mycangialbeetles, it is not known if fungi supplement the insectdiet [3, 5, 63]. A non-mycangial species, such as D.valens, requires an additional source of nitrogen. Thus, thebiological nitrogen fixation may provide part of thenitrogen demand required for growth, development, andreproduction. Nitrogen fixation in insect guts has beendemonstrated extensively in termites [47]. In this work,live adults and larvae of D. valens reduced acetylenemarkedly, several typical nitrogen-fixing bacteria wereisolated, and nifD genes were detected in the metagenomicDNA from guts. These evidences provide support for thenotion that bacterial biological nitrogen fixation occurs inD. valens gut. However, more studies must be performedto assess the proportion of nitrogen incorporated to thediet by symbiosis.

According to our results, we estimated that nitrogenequivalents of 5.6 and 4.4 µg of protein are fixed per larvaor adult per day. Nitrogen fixation is highly regulated at thetranscriptional level by a complicated regulatory networkthat responds to multiple environmental cues [28]. Thiscould explain the variability in the acetylene reductionassay (ARA). The possible roles performed by the bacterialcommunity associated with the gut of D. valens aresummarized in Fig. 5. Further studies on the bacterialcommunity are necessary to elucidate the nitrogen-fixingability in D. valens and to infer or support possiblefunctions of the different bacterial species detected in thisstudy.

Acknowledgments We thank Esperanza Martínez-Romero andMarco Rogel for their technical assistance with the acetylenereduction assay, and Francisco Bonilla, Marco Espinal, FernandaLópez, Arturo Vera, Javier Zavala for support in insect collection.Also, we thank Hugo Ramírez Saad and Félix Garrido for theirtechnical assistance with DGGE. This work was supported bygrants CGPI20060532 and CGPI20070651, IPN, and CONAFOR2002 C01-6020. Jesús Morales-Jiménez thanks to CONACyT andPIFI, IPN, for scholarships.

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