MicrobialEcology

Effect of Marine Bacterial Isolates on the Growth and Morphologyof Axenic Plantlets of the Green Alga Ulva linza

Katrina Marshall1,2, Ian Joint2, Maureen E. Callow1 and James A. Callow1

(1) School of Biosciences, University of Birmingham, Edgbaston, Birmingham B15 2TT, UK(2) Plymouth Marine Laboratory, Prospect Place, The Hoe, Plymouth PL1 3DH, UK

Received: 8 August 2005 / Accepted: 9 January 2006 / Online publication: 2 August 2006

Abstract

The green marine macroalga, Ulva linza, adopts anBatypical^ form when grown in the absence of bacteria.Twenty unique strains of periphytic bacteria, isolatedfrom three species of Ulva, were identified by 16S rDNAsequencing. These isolates were assessed for their effecton the growth and morphological development of axenicplantlets of U. linza. Results showed that the effect ofbacterial strains was strain- but not taxon-specific.Thirteen isolates returned the aberrant morphology tonormal and of these, five also significantly increasedgrowth rate. One isolate increased growth, but had noeffect on morphology. Biofilms of some of these isolatesstimulated the settlement of Ulva zoospores but therewas no correlation between bacterial isolates thatstimulated zoospore settlement and those that initiatedchanges in morphology and/or growth of the culturedalga.

Introduction

Epiphytic bacteria are abundant and ubiquitous colo-nizers of the external surfaces of marine macroalgae [2,18, 30], but relatively little is known about the nature oftheir interaction or the perceived benefits that followfrom the association. The bacteria are generally assumedto benefit through the ready availability of a range oforganic carbon sources produced by the alga [3]. Theadvantages to the alga are less obvious and although themechanism(s) is not well understood, it has been knownfor over 20 years that bacteria are involved in thedevelopment of normal morphology in green seaweeds.Provasoli and Pintner [26] showed that Ulva lactuca

plants had atypical Bpincushion^ morphology whengrown axenically. When these aberrant forms wereexposed to bacteria, the plants reverted to a normalBfoliaceous^ morphology. Similar effects on morphologyhave been observed when bacteria are excluded fromcultures of other green algal species, including U. pertusa[20, 21]; U. linza (syn. Enteromorpha linza) [9]; U.compressa (syn. E. compressa) [9]; and Monostromaoxyspermum [16, 34]. However, no study of this typehas combined the use of phylogenetically well-character-ized bacterial strains (i.e., those identified from 16SrDNA sequences) and a quantitative approach to assess-ing their effects on the growth and development of thealga.

A key question is the extent of specificity involved inthe interaction. Previous studies have investigated thisaspect with other species of green algae but the results areequivocal. Nakanishi et al. [20] reported that manybacterial genera, identified by using traditional microbi-ological techniques, were involved in promoting normalmorphogenesis of U. pertusa including Cytophaga,Flavobacterium, Vibrio, Pseudomonas, Halomonas, Escher-ichia, and Gram-positive cocci. On the other hand, it hasalso been suggested that morphogenesis in green macro-algae (Ulvaceae and Monostromaceae) is controlled by arestricted group of bacteria in the Bacteriodetes phylum,specifically Cytophaga and Flavobacterium spp. [16, 17,21].

In addition to influencing morphology and growththrough the periphytic microbiota, bacteria may haveother effects on algae, including their ability to colonizesurfaces. Patel et al. [25] showed that biofilms of specificstrains of marine bacteria influence the settlement ofzoospores of U. linza, and Joint et al. [13] showed thatone mechanism behind this may reside in the ability ofzoospores to detect the N-acylhomoserine lactones usedby many Gram-negative bacteria for quorum sensing.Correspondence to: James A. Callow; E-mail: j.a.callow@bham.ac.uk

DOI: 10.1007/s00248-006-9060-x & Volume 52, 302–310 (2006) & * Springer Science+Business Media, Inc. 2006302

Therefore, in this article we also test the hypothesis thatthere may be a relationship between those bacterialstrains that promote initial colonization by zoospores ofUlva, and those that are involved in promoting correctgrowth and development as periphytic strains.

Methods

Preparation of Axenic Algal Tissue. Vegetative andfertile sporophytic U. linza plants were collected betweenMarch and November in each year between 2001 and2004, from Wembury Beach, Devon, England (50-180N;4-020W). Plants were blotted to remove excess water, andstored on ice for transport back to the laboratory. Axeniccultures of Ulva were obtained by treating withantibiotics [28]. Loose debris was removed from thethallus, and pieces of the fertile apical region (approx. 1cm2) were placed in 50 mL artificial seawater (ASW)containing antibiotics that had been filter-sterilized usinga sterile 0.2-mm syringe filter. The antibiotic mix usedcomprised penicillin G (100 mg L

_1), streptomycin (25mg L

_1), norfloxacin (1 mg L_1), and kanamycin (25 mg

L_1). Germanium dioxide (1 g L

_1) was included toinhibit the growth of diatoms. Flasks containing treatedmaterial were maintained at 4-C in the dark for 4 days;this treatment maximized bacterial mortality whileminimizing any negative effects on the algae byreducing photosynthesis and lowering temperature.Thallus strips were subsequently washed withautoclaved ASW, placed in sterile ASW in 90-mm petridishes, and incubated at 18-C (irradiance 500 mmolquanta m

_2 s_1) until zoospores were released. Using

aseptic procedures, the zoospore mix was collected andspread onto an artificial algal medium [31] solidifiedwith 1.2% agar. Plates were incubated for 35 days in a14:10 h light/dark regime at 18-C. Axenic calli werepicked off and transferred to Stratmann’s medium [31].

Isolation of Bacteria. Bacteria were isolated fromfresh vegetative thallus of three species of Ulva (U. linza,U. compressa, and U. lactuca) using the method describedby Nakanishi et al. [20]. Material was gently washed inautoclaved ASW and 1 g of thallus tissue crushed in 10mL autoclaved ASW. After centrifugation at 3000 rpmfor 2 min to remove the larger cell debris, thesupernatant was serially diluted (100–10

_6) and spreadonto plates of four different media: oligotrophic agar [1],seawater plate count agar [24], M172 marine Cytophagamedium (http://www.dsmz.de/media/med172.htm), andseawater R2A [24, 32]. The plates were incubated at 20-Cfor 5 days, and individual colonies were picked off andstreaked onto the agar on which they were isolated toobtain single colonies. Isolation was carried out inautumn 2001 and bacterial isolates were maintained at_70-C in glycerol as source cultures for all experiments.

Fingerprinting Bacteria Isolates. Bacteria were ini-tially screened by denaturing gradient gel electrophoresis(DGGE) to remove replicates in the isolate collection.DNA was extracted from bacterial colonies using theCTAB method. Partial 16S rDNA was amplified usingpolymerase chain reaction (PCR). The variable V3 regionof the 16S rDNA from members of the domain Bacteriawas amplified using the PRBA338F primer (50-ACTCC-TACGGGAGGCAGCAG-30; Escherichia coli positions338–358) [22] and the PRUN518R primer (50-ATTACCGCGGCTGCTGG-30; E. coli positions 534–518) with a GC clamp [19]. The reaction mixturecontained 1� PCR buffer, 3 mM MgCl2, 100 ng of eachforward and reverse primer, 20 mM of each deoxynu-cleoside triphosphate (dATP, dCTP, dGTP, dTTP), 1 mgmL

_1 bovine serum albumin (BSA), 1 unit of Taqpolymerase, and õ10 ng of template DNA. The PCRprotocol included a 5-min initial denaturation at 95-C,followed by 30 cycles at 94-C for 30 s, 55-C for 30 s,72-C for 30 s, with a final cycle of 10 min at 72-C andincubation at 4-C until processed further. DGGE wasundertaken using a D-Gene Mutation Detection System(Bio-Rad, USA). The gel was 8% (w/v) polyacrylamide(37.5:1 acrylamide/bisacrylamide) in 0.5 M Tris–acetate–EDTA (TAE) with a denaturant gradient between 30 and60% [100% denaturant contained 7 M urea and 40% (v/v) formamide in 0.5� TAE]. Gels were run at 40 V for 16h at 60-C, removed, and stained with ethidium bromide(5 mL in 50 mL 1� TAE) for 1 h, and subsequentlydestained in 1�TAE for 10 min before visualization witha UV transilluminator.

Phylogeny of 16S rDNA. PCR amplification of thepartial 16S rDNA from single isolates used the universalforward bacterial primer pA (50-AGAGTTT GATCCTGGCTCAG-30; E. coli positions 8–27) and the reverseprimer pH (50-AAGGAGGTGATCCAGCCGCA-30; E.coli positions 1541–1522) [15]. The reaction mixturecontained 1� PCR buffer, 3 mM MgCl2, 100 ng each offorward and reverse primer, 20 mM of eachdeoxynucleoside triphosphate (dATP, dCTP, dGTP,dTTP), 1 unit of Taq polymerase, and õ10 ng oftemplate DNA. The protocol included a 3-min initialdenaturation at 95-C, followed by 30 cycles at 95-C for 30s, 55-C for 30 s, 72-C for 45 s, finally 1 cycle of 10 min at72-C and storage at 4-C. Product was run on a 1% lowmelting point agarose gel at 40 V and the bands wereexcised and purified using QIAquick Gel Extraction kit(Qiagen, no. 28704). The purified product wasresuspended in sterile water treated with diethylpyrocarbonate (DEPC). Sequencing of isolates was bythe Genomics Laboratory at the University ofBirmingham. A search of the EMBL databasewww.ebi.ac.uk/blast2/nucleotide.html) for similarity of16S rDNA sequences to isolated strains was made using

K. MARSHALL ET AL.: EFFECT OF BACTERIA ON THE ALGA ULVA LINZA 303

the WU-BLAST2 program [10]. All isolate sequences andother related marine strains were aligned usingCLUSTAL W [35]. The 16S sequences of the isolatedstrains have been submitted to Genbank with theaccession numbers AM180731–AM180750.

Assessing the Effect of Bacteria on Morphogenesis

and Growth of U. linza. A nondestructive method wasrequired to estimate algal growth and procedures weredeveloped based on image analysis of algal surface area.Batches of four axenic plants of similar size weretransferred to 12-well plates containing 3 mL Stratmannmedium. The influence of each of 20 bacterial isolates ongrowth and morphology was assessed in experiments withfour replicates of each isolate. Plants were incubated at18-C in a 14:10 h light/dark cycle (irradiance, 500 mmolquanta m

_2 s_1) for 28 days after inoculation with

bacteria. Media were refreshed and bacteria reinoculatedevery 7 days. Digital images of plants were taken every 7days and plant area was used as a proxy for growth, thepixel area of each plant image being determined with ZeissKS300 image analysis software. Growth of U. linza plantswas expressed as the Bcumulative relative growth rate^(RGR) over the 28-day period, determined as percentageincrease in area per day [14]. Data were tested fornormality via Andersen–Darling test [23] andhomogeneity of variance using Minitab 13.31. The dataconformed to both and were analyzed using a one-wayANOVA and Dunnett’s family post-hoc comparison.

Plant morphology was assessed on a semiquantitativescale based on the number and state of development oftubules extending from the central callus of each plant.The greatest difference in morphology compared toaxenic controls was assigned a score of 3; in practice,this indicated the growth of 950 well-developed tubularextensions. Intermediate levels of change were assignedscores of 2 and 1, with between 30–50 and 10–30 tubularextensions from the central callus, respectively. Thesmallest difference compared to the control, with growthof 0–10 tubular extensions, was assigned 0.

Direct Observation of Bacteria on Ulva sur-

faces. The presence of bacteria on the plant surfaceswas determined after 28 days. Plants were stained with0.001% (w/v) filter-sterilized acridine orange for 45 s inthe dark, washed gently 3� with sterile distilled waterand observed under epifluorescence using an Axioskop 2(Zeiss) microscope and an FS 10 (FITC) filter. Numbersof bacteria on plant surfaces were estimated semiquan-titatively by using �40 objective. The numbers ofbacteria in five focal planes were summed and scoredusing the following scale based on the number of bacteriavisible on a plant area of 0.001 mm2: (0) no bacteriavisible, (1) 1–100 bacteria (2) 101–200 bacteria, (3) 201–300 bacteria, (4) 9300 bacteria.

Effect of Bacterial Isolates on Zoospore Settle-

ment. Each of the 20 bacterial isolates was investigat-ed for its ability to attract zoospores by using the methoddescribed by Joint et al. [12]. Bacterial biofilms were de-veloped on glass coverslips by inoculating with 7.5 mL ofstationary phase culture of individual bacteria grown inmarine broth, and incubated at room temperature for 72h. Three coverslips were placed into each of two sterile 90-mm diameter petri dishes, one acting as a control that wasnot exposed to zoospores and the other treated with zo-ospore suspension. Zoospores were released from fertilesporophytic thallus using the method described by Callowet al. [6]. The zoospore concentration was adjusted to 1�106 zoospores mL

_1, and added to three replicate treat-ments as described by Joint et al. [12]. The proportion ofbacterial surface area that was covered by zoospores wascalculated, and the stimulation of zoospore settlement byeach bacterial strain was estimated [25].

The data were tested for normality and homogeneityof variance using Minitab 13.31. One-way ANOVA and apost-hoc Tukey’s B test were used for the zoospore andzoospore-adjusted settlement as described in Patel et al.[25]. Data for preferential settlement were compared byusing a two-sample t test. The degree of settlement wascalculated by using the critical values after Tukey’s B test(for zoospore and zoospore-adjusted) and t test (forBpreferential^), whether 1�, 2�, 3�, or 4� greater thanthe test statistics, respectively [27]. Correlations betweengrowth, morphology, and settlement data were calculatedby using Pearson correlation (Minitab 13.31).

Results

U. linza in its normal, intertidal habitat displays amonostromic, tubular morphology, i.e., a hollow tubularthallus one cell thick. The aberrant morphology thatdevelops in axenic plantlets of U. linza is shown in Fig. 1.In the absence of bacteria, tubule formation is inhibitedand the plant forms a compact undifferentiated callus ofcells without distinct morphology (Fig. 1a). In contrast,cultivated plantlets grown in the presence of bacteria (inthis case the natural periphytic consortium) producenumerous tubular structures, each of which is capable ofdeveloping into a mature thallus.

Effect of Individual Isolates on Growth and

Morphology. The collection of 38 unique isolates wasscreened for their effects on growth and morphology ofaxenic plantlets in a preliminary experiment. From this, asubset of 20 strains was selected for more detailedexperiments. The subset of isolates included thosesequenceable strains that gave a clear effect in thepreliminary experiment plus a small number that hadno effect, as controls. Some bacteria had a markedinfluence on plantlet growth rate. After 28 days, the total

304 K. MARSHALL ET AL.: EFFECT OF BACTERIA ON THE ALGA ULVA LINZA

relative increase in plant area after inoculating withdifferent isolates was between 3% (UL30) and 243%(UC15) greater than control, axenic plants (Table 1, Fig.2). The majority of isolates [14] produced only smallincreases in growth rate (between 20 and 100%), whichwere not statistically different to the controls. However,six isolates (ULA2, ULA5, UL16, UC15, UC19, UC38)significantly increased relative growth rates (F=4.14,pG 0.001) compared to the control treatment plants(Fig. 2).

There was little difference in the type of morphologyinduced by the individual bacterial isolates. All effectiveisolates induced the formation of tubular extensionsfrom the central callus-like tissue. However, there weredistinct differences between isolates in the extent ofmorphological change induced over the 28-day experi-mental period (Table 1). Examples of morphologiesobserved after 28 days are shown in Fig. 1c–f. Control,uninoculated plants were callus-like in form (Fig. 1c).

Three isolates did not cause any marked change inplantlet morphology (i.e., scores of G1) (Table 1). Onebacterial isolate (UC38) did not cause a change inmorphology, although it did induce a significant increasein growth. Thirteen isolates stimulated a marked mor-phological change with over 30 well-developed tubularextensions (morphological index Q2) (Table 1). Five ofthese (ULA2, ULA5, UL16, UC15, UC19) were repre-sented in the six strains that also caused a significantincrease in the RGR.

Bacterial Density on Plant Surfaces. Epifluores-cence microscopy confirmed that control plants re-mained axenic because no bacteria were observed onplant surfaces when stained with acridine orange. Onplants that developed normal morphology, the distribu-tion of bacteria was patchy, with generally fewer bacteriapresent on the tubular extensions than the rest of theplant. In practice, it was easier to see bacteria on thetubules than in the dense tissues at the center of the plantwhere is was difficult to accurately quantify bacterialnumbers due to high absorption of stain by the plant cellsand because of the density of plant tissue requiredfrequent alteration of the focal plane.

Bacterial density was assessed on a semiquantitativescale (Table 1). Three isolates (ULA16, UC15, UC19)grew to densities 92 � 105 bacteria mm

_2 and all ini-tiated marked changes in plant morphology (morpho-logical index Q2, Table 1). Two of these isolates (UC15,UC19) also significantly enhanced the growth of plant-lets. Three isolates (ULA1, UL2, UL5) could not be ob-served on the plant surface, but resulted in somemorphological change (index G2); none had a significanteffect on plant growth. Of the remaining 14 isolates, thenumbers on the plant surfaces were much lower, rangingfrom 1 � 102 to 2 � 105 bacteria mm

_2. One of theseisolates (UC38) significantly stimulated growth butcaused little morphological change, whereas 10 initiatedthe production of 930 tubular extensions; three isolates(ULA2, ULA5, UL16) also significantly stimulatedgrowth.

In general terms, there was little correlation betweenthe numbers of bacteria present on the plantlet surfacesand either changes in morphology or growth of Ulva.Some strains that flourished on the plant surface(UC15, UC19) had significant effects on growth andstrong effects on morphology, whereas one strain (ULA16)that reached high surface abundance (score 4) did notsignificantly increase growth and had only a modesteffect (score 2) on morphology. Some strains (UIA1,UL2, UL5) clearly did not flourish on the plantletsurface with no recorded abundance in direct micro-scopic evaluation. None of these had a significant ef-fect on plant growth and, at best, had only a minoreffect on plant morphology. On the other hand, some

Figure 1. Morphologies of plantlets of Ulva linza cultured fromreleased spores. (a) Example of a 6-week-old axenic plantletcultured from spores that were released from antibiotic-treatedplants. (b) Portion of a 6-week-old cultured, but nonaxenicplantlet derived from spores released from plants that had notbeen pretreated with antibiotics, showing the characteristic tubularmorphology. (c)–(f ) Representative morphologies of U. linzaplants 28 days after inoculation with bacterial isolates: (c)control—no bacteria (morphology score 0); (d) inoculated withisolate UL4 (morphological score 1); (e) inoculated with isolateUL19 (morphological score 2); (f ) inoculated with isolate UC19(morphological score 3). All scale bars = 1 mm.

K. MARSHALL ET AL.: EFFECT OF BACTERIA ON THE ALGA ULVA LINZA 305

strains (ULA5, UL16, UC38) had modest abundance onthe plant surface (score 1), yet all three had statisticallysignificant effects on growth, and in two cases an effect onmorphology. One strain (ULA16) had high surfaceabundance but did not significantly stimulate growth,and had only a moderate (score 2) effect on morphology.

Phylogenet ic Character izat ion of Bacter ia l

Isolates. Bacteria used in these experiments wereisolated from the surfaces of three species of Ulva. Intotal, 106 strains were isolated, 40 from U. compressa, 35from U. linza, and 31 from U. lactuca. After reduction ofredundancy using DGGE fingerprinting, 38 uniqueisolates were obtained. Eight of these were Gram-positive (although none of these was isolated from U.linza). Partial 16S rDNA sequences (approx. 1400 bp) ofthe bacterial isolates were amplified and sequenced. Theclosest homologous sequences in the EMBL database areshown for the 20 isolates that were selected from theoriginal collection of 38, for detailed studies on growth

and morphology of Ulva plantlets (Table 1). The highestproportion of strains was found within the Proteobac-teria and the Bacteriodetes phyla.

Effect of Bacteria on Zoospore Settlement. Sporesettlement experiments were carried out with biofilms ofthe bacterial isolates to ascertain whether there was arelationship between settlement-modifying strains andthose that influence growth and morphology.Interestingly, none of the isolates inhibited settlement ofUlva zoospores—which would certainly be incompatiblewith a role in controlling morphology—but significantdifferences in the ability of isolates to enhance zoosporesettlement were detected.

Increased settlement was assessed by comparing thenumbers of attached zoospores on a clean surface withthe number on the same surface supporting a bacterialbiofilm. In addition, the spatial relationship betweenzoospores and bacteria was investigated. The density ofbacteria on control slides (not exposed to zoospores)

Table 1. Influence of 20 bacterial isolates on morphology and growth rate of Ulva linza plants after 28 days incubation

Isolate IDRGR

(% day_1)a

Morphologyscoreb

Bacterial abundancescorec

Closest matchingstrain in EMBL Phylum

Accessionnumber

% Sequencesimilarity

Control 5.57 T 0.59 –ULA1 6.17 T 0.67 0–1 0 Frigoribacterium sp. Actinobacteria AF157478 98ULA2 7.31 T 0.95* 2–3 2 Paracoccus

zeaxanthinifaciensProteobacteria

(alpha)AF461159 94

ULA5 7.73 T 1.27* 2 1 Psychrobacter fozii Proteobacteria(gamma)

AJ430827 95

ULA10 6.51 T 1.15 2 2 Uncultured Bacillus sp. Firmicutes AY082370 95ULA16 6.42 T 1.41 2 4 Cellulophaga lytica Bacteroidetes AB032512 97ULA23 6.53 T 1.28 2–3 2 Ruegeria sp. Proteobacteria

(alpha)AY005463 91

UL2 5.46 T 1.34 1–2 0 Bacteroidetes bacterium Bacteroidetes AF539760 94UL3 6.08 T 1.25 2–3 1 Rhodobacter apigmentum Proteobacteria

(alpha)AF035433 95

UL4 4.25 T 1.55 1 2 Marine bacteriumisolated fromgreen alga

AF536383 93

UL5 4.91 T 0.97 1–2 0 Gelidibacter sp. Bacteroidetes AF513399 93UL16 7.34 T 0.76* 3 1 Cellulophaga sp. Bacteroidetes AF539757 92UL19 5.81 T 0.92 2 2 Shewanella gaetbuli Proteobacteria

(gamma)AY190533 96

UL30 5.13 T 0.82 0–1 1 Cytophaga sp. Bacteroidetes AF235126 95UL34 6.79 T 0.78 2 1 Pseudoalteromonas citrea Proteobacteria

(gamma)AF529062 99

UC14a 6.49 T 1.07 1–2 2 Pseudomonas sp. Proteobacteria(gamma)

AF500211 97

UC15 8.60 T 1.17* 3 3 Glacial ice bacterium AF479366 89UC19 8.53 T 0.75* 3 3 Cytophaga sp. Bacteroidetes AB073591 98UC21 7.24 T 0.99 3 2 Planococcus citreus Firmicutes AF500008 98UC28 6.95 T 1.01 2 1 Cobetia marina Proteobacteria

(gamma)AJ306890 97

UC38 7.98 T 0.74* 0 1 Planococcus maritimus Firmicutes AY428552 98aAverage relative growth rate (RGR, percentage increase in area per day, n = 16) T 95% confidence interval. Isolates marked by an asterisk (*) showedsignificant increases (pG0.05) in RGR of plants compared to the control using one-way ANOVA and Dunnett’s post-hoc analysis.bMorphology after 28 days assessed on a semiquantitative scale. 0: Little tubular growth (G10) from central callus; 1: 10–30 tubular extensions; 2:30–50 tubules; 3: 950 well-developed tubules.cBacterial abundance based on total numbers of bacteria from five focal planes in 0.001 mm2 using a �40 objective. 0: No visible bacteria; 1: 1–100 bacteria;2: 101–200 bacteria; 3: 201–300 bacteria; 4: 9300 bacteria. Accession numbers shown are those of the closest match in the EMBL–EBI database.

306 K. MARSHALL ET AL.: EFFECT OF BACTERIA ON THE ALGA ULVA LINZA

was compared with the density of bacteria visible onslides exposed to zoospores. If similar bacterial densitieswere observed, zoospore settlement would be consid-ered random. A positive or negative difference forthe expected distribution of visible bacteria results whenzoospores settle on or away from the bacterial cells,respectively.

Nine isolates significantly stimulated zoospore set-tlement (Fig. 3). Four of these also effected a markedchange in the plant morphology (ULA5, ULA10, UL16,and UC28), whereas two enhanced growth rate (ULA5,UL16). In addition, five isolates (ULA5, UL2, UL34,UC19, and UC28) showed highly preferential settlementof the zoospores directly onto bacterial cells at levels 3

and 4 times higher (critical t values) than the teststatistic. Three of these (ULA5, UC19, and UC28) causeda substantial change in plant morphology (morphologyindex Q2) and two (ULA5, UC19) a significant increasein growth of the U. linza plantlets. Of the seven strainsthat did not stimulate preferential settlement of zoo-spores onto bacterial cells, two (ULA10, UL3) stimulatedmorphological changes in the U. linza plants, one(UC38) stimulated a significantly increased RGR, andtwo (ULA2, UL16) stimulated both changes in morphol-ogy and increased RGR.

Although individual strains showed both stimulatoryeffects on zoospore settlement and preferential settle-ment, there was no overall correlation between the ability

0

200

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ULA

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ULA

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ULA

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16

ULA

23

UL2

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UC

14a

UC

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UC

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UC

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Bacterial isolate

% S

timul

atio

n

*+

*#+

*#+

*++

* +

+

++

+

#+

#+

#

#

#

#

#

Figure 3. Percentage stimulation ofU. linza zoospore settlement onglass coverslips with 72 h mono-species bacterial biofilms. Barsare mean T95% confidence interval(n=60). #Values significantly differ-ent to controls. *Isolates that stim-ulated a significant increase ingrowth of U. linza plants. +Isolatesthat stimulated a marked change inmorphology (930 tubules) in U.linza plants.

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trol

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Isolate

Rel

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e in

crea

se in

are

a(m

m2m

m-2)

** *

**

*

Figure 2. Relative increase in sur-face area of U. linza plants after 28days of incubation with 20 individ-ual strains of bacteria. Controlexperiments were axenic plantswithout added bacteria. Bars repre-sent mean T 95% confidence interval(n=16). *Isolates with a significantlydifferent relative growth rate from

K. MARSHALL ET AL.: EFFECT OF BACTERIA ON THE ALGA ULVA LINZA 307

to simulate Ulva zoospores settlement with eitherstimulation of plantlet growth rate and/or morphologicalchanges in the plant.

Discussion

This study has demonstrated that the morphology andgrowth rate of U. linza are influenced by bacteria, andthat Bnormal,^ wild-type morphology is absolutelydependent on the presence of bacteria. These findingsare consistent with previous observations on axenic Ulvaspp. (syn. Enteromorpha spp.) [9] and U. lactuca [26].However, previous studies were not performed withstrains of bacteria that were phylogenetically wellcharacterized, so the question of specificity in thebacterium–alga interaction could not be fully addressed.In the present study, using isolates identified from their16S sequences, it appears that there is some specificitybecause from a total of 38 isolates only 6 caused astatistically significant effect on plant growth. Rather,more isolates caused a distinct effect on plantletmorphology but at least 15 of the original 38 isolateshad no effect on either growth or morphology. Withinthe group of effective isolates there is also somespecificity in terms of the degree of the effects observed,and in one interesting case (UC38, 98% similar toPlanococcus maritimus at the 16S rDNA level) growthwas affected without a change in the morphology.However, there is clearly no taxon specificity in theeffects observed because the effective isolates are distrib-uted over several bacterial taxa. Nakanishi et al. [20] havesimilarly reported that many bacterial genera are in-volved in morphogenesis of U. pertusa including Cyto-phaga, Flavobacterium, Vibrio, Pseudomonas, Halomonas,Escherichia, and Gram-positive cocci. On the other hand,it has also been suggested that morphogenesis in greenmacroalgae (Ulvaceae and Monostromaceae) is con-trolled by a restricted group of bacteria, specificallyCytophaga and Flavobacterium spp. [17, 21]. In contrast,in the present study, only three of the seven isolates fromthe Bacteriodetes group initiated a change in morphol-ogy of U. linza plants. A number of isolates have notpreviously been reported to have an effect on morphol-ogy or growth; these include isolates with close 16S ho-mology to Psychrobacter sp. (ULA5), Pseudoalteromonassp. (UL34), and Shewanella sp. (UL19) in the gammap-roteobacteria. Both Pseudoalteromonas sp. and Shewanellasp. are known to influence the settlement of Ulva zoo-spores, with some isolates inhibiting and others enhanc-ing settlement on unibacterial biofilms [25]. In thepresent study, none of the isolates from these generastimulated zoospore settlement. Gram-positive bacteriawere also found to affect the morphology or growth of U.linza to a greater extent than reported by Nakanishi et al.[20] for U. pertusa.

The growth rate of other marine algae has beenshown to be affected by bacteria. The toxic dinoflagellateGambierdiscus toxicus increased growth rate by 65.4%when axenic cultures were inoculated with bacteria [29].However, the effect of bacteria on the growth rate ofmacroalgae has not been previously quantified, althoughsubjective indications of an acceleration in growth anddevelopment have been reported [17] for sporelings of U.pertusa, U. conglobata, and U. intestinalis when inoculat-ed and incubated with strains of bacteria of theBacteriodetes phylum.

The mechanism by which bacteria modulate themorphology of the plant is not yet understood, althougha number of hypotheses have been suggested. Anendosymbiotic bacterium from the Agrobacterium–Rhi-zobium group, containing the nifH gene encoding fornitrogenase, was isolated from rhizoids of the green alga,Caulerpa taxifolia [7]. It was suggested that this strainmight be important for nitrogen supply to the seaweed.In the red alga, Prionitis lanceolata, gall formation isassociated with a bacterium of the Roseobacter group. Inthe galls, there is overproduction of indole-3-acetic acid,but the role of this, and the bacterium upon thephysiology of the alga is as yet undetermined [5]. Ifbacteria are involved in the production or turnover ofsuch plant hormones, this may be one mechanism bywhich bacteria influence morphology and increasedgrowth rate of U. linza plants [5]. It has also beensuggested that secondary metabolites released by someepibiotic bacteria may prevent subsequent biofouling byother organisms [4, 11], thereby providing some protec-tion to the host alga. As yet, these are few data to supportthese speculations.

It is not clear if contact between bacteria and plants isnecessary for morphology to be affected. U. pertusa isreported to require physical contact with species ofCytophaga and Flavobacterium [21] because mutantstrains that could not attach to the algae did not inducemorphogenesis. In the present study, the degree ofattachment of the bacterial isolates varied and noparticular pattern was detected. The numbers attachedwere comparable to other studies [8], but distribution waspatchy on the plantlet surface. Some isolates that inducedmarked morphology and growth changes (e.g., UL16)had very low bacterial numbers attached to the plants.This suggests either that direct contact is not essential orthat morphology can be changed by the presence of verylow densities of bacteria. Alternatively, extracellularsubstances produced by bacteria might initiate change,and attached bacteria may not be necessary. In otherstudies, supernatants have been shown to be effective. Inthe case of M. oxyspermum, extracts of brown and redalgae [34] and supernatants of bacterial cultures [16, 34]have restored the natural morphology of the alga. It hasbeen proposed by Matsuo et al. [17] that an exogenous

308 K. MARSHALL ET AL.: EFFECT OF BACTERIA ON THE ALGA ULVA LINZA

growth factor, identified as thallusin, produced bybacteria belonging to the Bacteriodetes grouping, is anessential factor for normal growth of M. oxyspermum.However, in work undertaken by the authors of thisstudy, it was found that supernatants from bacterialstrains other than the Bacteriodetes also stimulated thegrowth and morphology of U. linza (unpublished data).

No overall correlation was found between isolatesthat altered the morphology or growth of U. linza plantsand those that enhanced zoospore settlement. Somestrains affected both settlement and plant morphology;e.g., isolate UC15 stimulated both settlement andmorphology. In general, isolates that had the greatesteffect on zoospore settlement did not show any overallcorrelation with the initiation of changes in morphologyor influence growth rate of axenic plants. However, threeisolates (ULA5, UC15, and UC19) stimulated zoosporesto settle and initiated changes in both growth andmorphology in the axenic plants.

A range of different bacteria in this study stimulatedchanges in morphology or growth of Ulva, suggestingsome redundancy in ecological function. That is, normalmorphology is not dependent on the presence of a uniquebacterium, but differentiation can be effected by a rangeof different bacteria. However, it should be rememberedthat in this study we only considered those bacteria thatwere cultivatable. It is well known that only a very smallproportion of marine bacteria can be cultivated bycurrent methodologies, so the results presented here onlyapply to a small proportion of bacteria that are on thesurface of a seaweed or in the water column. Anundetermined number of bacterial species may have theability to influence algal morphology.

In summary, this article has confirmed that bacteriaare essential for the normal growth and development ofthe alga U. linza. Several different genera of bacteriainfluence the growth and morphology of the alga. Theeffect of bacteria on the plants is not taxon-specific andcannot be assigned to a species or a genus. The widerange of bacteria that initiate changes in the morphologyor growth presumably confers ecological flexibility on thealga so that it is not dependent on a small number ofspecific bacteria. This may be important when thepresence of different bacteria may depend on factorssuch as season, temperature, or desiccation in intertidalregions. The mechanisms involved in these bacterial–algal interactions are not known, but direct attachmentof bacteria to the plant does not appear to be essential.The morphogenic switch occurs either through theattachment of very sparse bacterial assemblages or byproduction of exogenous factors by bacteria. Finally,there is no strong evidence that the bacteria that areimportant for attracting zoospores to surfaces [13, 33]are the same bacteria that are necessary for the normaldevelopment of Ulva plants.

Acknowledgments

This research was support by the Natural EnvironmentResearch Council through grant number NER/T/S/2000/00623. We thank Karen Tait for helpful advice onbacterial cell signaling.

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