Polyphasic assessment of fresh-water benthic mat-forming cyanobacteria isolated from New Zealand

R E S E A R C H A R T I C L E

Polyphasic assessmentoffresh-water benthicmat-formingcyanobacteria isolated fromNewZealandMark W. Heath1, Susanna A. Wood2 & Ken G. Ryan1

1School of Biological Sciences, Victoria University of Wellington, Wellington, New Zealand; and 2Cawthron Institute, Nelson, New Zealand

Correspondence: Susanna A. Wood,

Cawthron Institute, Private Bag 2, Nelson

7001, New Zealand. Tel.: 164 3 548 2319;

fax: 164 3 546 9464; e-mail:

[email protected]

Received 26 October 2009; revised 25 February

2010; accepted 2 March 2010.

Final version published online 4 May 2010.

DOI:10.1111/j.1574-6941.2010.00867.x

Editor: Riks Laanbroek

Keywords

anatoxin-a; benthic cyanobacteria;

Phormidium; Phormidium autumnale.

Abstract

Mat-forming benthic cyanobacteria are widespread throughout New Zealand

rivers, and their ingestion has been linked to animal poisonings. In this study,

potentially toxic benthic cyanobacterial proliferations were collected from 21 rivers

and lakes throughout New Zealand. Each environmental sample was screened for

anatoxins using liquid chromatography-MS (LC-MS). Thirty-six cyanobacterial

strains were isolated and cultured from these samples. A polyphasic approach was

used to identify each isolate; this included genotypic analyses [16S rRNA gene

sequences and intergenic spacer (ITS)] and morphological characterization. Each

culture was analysed for anatoxins using LC-MS and screened for microcystin

production potential using targeted PCR. The morphospecies Phormidium

autumnale was found to be the dominant cyanobacterium in mat samples.

Polyphasic analyses revealed multiple slight morphological variants within the

P. autumnale clade and highlighted the difficulties in identifying Oscillatoriaceae.

Only one morphospecies (comprising the two strains CYN52 and CYN53) of

P. autumnale was found to produce anatoxins. These strains formed their own

clade based on partial 16S rRNA gene sequences. These data indicate that benthic

P. autumnale mats are composed of multiple morphospecies and toxin production

is dependent on the presence of toxin-producing genotypes. Further cyanobacteria

are also characterized, including Phormidium murrayi, which was identified for the

first time outside of Antarctica.

Introduction

The first report of toxin production in planktonic cyanobac-

teria was published in 1878 (Francis, 1878). Since then,

multiple incidents of animal and human poisonings have

been linked to planktonic cyanobacteria and the cyanotoxins

responsible have been identified (Sivonen & Jones, 1999). In

contrast, there has been little information available on toxic

benthic cyanobacteria. However, over the past two decades,

an increasing number of toxin-producing freshwater benthic

cyanobacteria species have been documented (Edwards et al.,

1992; Baker et al., 2001; Mohamed et al., 2006; Fiore et al.,

2009), and ingestion of these has been associated with animal

poisonings (Hamill, 2001; Krienitz et al., 2003; Gugger et al.,

2005). In New Zealand, reports of dog poisonings linked to

benthic cyanobacteria have increased in the last 10 years.

Since the first dog fatalities were documented in 1998

(Hamill, 2001), there have been 4 30 reported deaths

(Wood et al., 2007, S.A. Wood, unpublished data.).

Globally, anatoxins (neurotoxins) and microcystins

(hepatotoxins) are the two most commonly produced cya-

notoxins by benthic cyanobacteria (Mez et al., 1997; Sivonen

& Jones, 1999; Cadel-Six et al., 2007; Izaguirre et al., 2007;

Wood et al., 2007). Anatoxins are powerful neuromuscular-

blocking agents that act through the nicotinic acetylcholine

receptor, while microcystins inhibit protein phosphatases

causing liver necrosis (MacKintosh et al., 1990; Carmichael,

1994). Recently, benthic cyanobacteria that produce cylin-

drospermopsins and saxitoxins have also been identified

(Carmichael et al., 1997; Seifert et al., 2007). Saxitoxins are

fast-acting neurotoxins that inhibit nerve conduction by

blocking sodium channels; these toxins are common in

marine dinoflagellates, where they are known as paralytic

shellfish poisons, (Adelman et al., 1982). Cylindrospermop-

sins are potent inhibitors of protein and glutathione synth-

esis acting on the liver and kidneys (Terao et al., 1994;

Falconer et al., 1999). There have been no reported cases of

animal toxicosis from benthic cyanobacteria producing

FEMS Microbiol Ecol 73 (2010) 95–109 c� 2010 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

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saxitoxins and cylindrospermopsins. Benthic Phormidium

and Oscillatoria sp. have most commonly been linked to

cyanotoxin production (Krienitz et al., 2003; Gugger et al.,

2005; Cadel-Six et al., 2007; Izaguirre et al., 2007; Wood et al.,

2007). However, cyanotoxins have also been found in benthic

species of Spirulina (microcystins and anatoxins), Fischerella

(microcystins), Lyngbya (saxitoxins and cylindrospermop-

sins), Aphanothece (microcystins) and Nostoc (microcystins)

(Carmichael et al., 1997; Krienitz et al., 2003; Dasey et al.,

2005; Seifert et al., 2007; Fiore et al., 2009).

In New Zealand rivers, benthic mat-forming cyanobac-

teria are found under a wide range of water quality

conditions (Biggs & Kilroy, 2000). The most common

mat-forming genus in New Zealand is Phormidium (Minis-

try for the Environment and Ministry for Health, 2009).

Under optimal conditions, Phormidium forms expansive

black/brown/green leathery mats over wide areas of river

substrate. In the 2005/06 summer, Wood et al. (2007)

identified the causative cyanobacterium of multiple dog

deaths in the Hutt River (Wellington, New Zealand) as

Phormidium autumnale. This organism is the only benthic

species known to produce anatoxin-a (ATX) and homoana-

toxin-a (HTX) in New Zealand. Routine testing of Phormi-

dium mats from around New Zealand has shown marked

variations in the presence of anatoxins, in the anatoxin

variants produced and in their concentrations. There is

uncertainty as to whether this variability is caused by the

presence of different strains within the mats or to variations

in their ability to produce anatoxins, or by environmental

triggers, i.e. temperature. The correct and early identifica-

tion of cyanobacterial species and confirmation of those

species that produce toxins will provide guidance that can be

used in developing management and mitigation pro-

grammes aimed at protecting animal and human health.

In this study, 31 potentially toxic benthic proliferations of

cyanobacteria were collected from 21 rivers and lakes around

New Zealand. Each environmental sample was screened for

anatoxins using liquid chromatography-MS (LC-MS). Indi-

vidual isolates from each sample were cultured and anatoxins

were analysed using LC-MS. Each culture was screened for

microcystin production potential using PCR. Isolates from

the pure cultures were identified where possible to the species

level using morphology and phylogenetic analyses. This

study is the first to describe the diversity and toxin produc-

tion of benthic cyanobacteria in New Zealand.

Materials and methods

Site description and sample collection

Between 2005 and 2008, benthic cyanobacterial mats were

collected from 21 New Zealand rivers and lakes experiencing

cyanobacterial proliferations. Sampling sites from which

cyanobacterial strains were successfully isolated are shown

in Fig. 1. Cyanobacterial mats were predominantly found on

rocky substrates, but were also collected from fine substrate

(0.2–0.02 mm) in the Whakatikei and Rangitaiki rivers.

Samples collected from the Waikato river were from a small

geothermal tributary. Samples were collected by scraping

mats into sterile plastic screw-cap bottles (50 mL, Biolab,

New Zealand). All samples were placed on ice for transport.

On arrival at the laboratory, samples were frozen immedi-

ately without cryopreservation (� 20 1C) for later culturing

and toxin analysis. Subsamples (10 mL) were preserved

using Lugol’s Iodine for morphological identification.

Strain isolation and culture conditions

Frozen cyanobacteria samples were thawed and cyanobac-

terial strains were isolated by streaking on a solid MLA

medium (Bolch & Blackburn, 1996). One half of the Petri

dish containing the streaked material was covered in black

PVC. This helped in isolating single filaments as some

benthic cyanobacteria are motile and move towards light.

After approximately 2 weeks, when filaments had moved

across the Petri dishes, single filaments were isolated

by micropipetting and transferred to 24-well plates contain-

ing 500 mL MLA medium per well. Filaments were

washed repeatedly and incubated under standard

conditions (100� 20mmol photons m�2 s�1; 16 : 8 h light : -

dark; 18� 1 1C, Contherm, 190 RHS, New Zealand). Cyclo-

heximide (100mg mL�1) was used in selected cultures to

reduce eukaryotic growth (Ferris & Hirsch, 1991; Urmeneta

et al., 2003). Successfully isolated strains were maintained in

50-mL plastic bottles (Biolab) under the above conditions.

Morphological identification

Subsamples of each cyanobacterial strain (in stationary

phase) were identified by microscopy (Zeiss Photomicro-

scope II, Germany). Photomicrographs were taken using a

digital camera (Canon Powershot S3IS) and further pro-

cessed in PHOTOSHOP 7.0 (Adobe). Nomarski interference

contrast microscopy was used in addition to bright-field

microscopy to enhance the contrast in unstained samples.

Species identifications were made primarily by reference to

Komarek & Anagnostidis (2005) and McGregor (2007).

Thirty measurements were made of vegetative cell lengths

and widths for each isolate, and detailed observations of

phenotypic characteristics were noted for 15 filaments.

Isolation of DNA and molecular characterization

Subsamples (500mL) from each culture were centrifuged

using an Eppendorf microcentrifuge (15 000 g, 1 min) and

the supernatant was removed by sterile pipetting. DNA was

extracted from the pellets using a PureLinkTM Genomic

FEMS Microbiol Ecol 73 (2010) 95–109c� 2010 Federation of European Microbiological SocietiesPublished by Blackwell Publishing Ltd. All rights reserved

96 M.W. Heath et al.

DNA kit (Invitrogen, CA) according to the Gram-negative

bacteria protocol supplied by the manufacturer.

PCR amplification of a segment of the 16S rRNA gene and

intergenic spacer (ITS) region were performed in 50mL

reaction volume containing between 100 and 200 ng of DNA,

0.5mM of each primer [16S rRNA gene used 27F and 809R

Jungblut & Neilan (2005), ITS used 322 and 340 Iteman et al.

(2000); Geneworks, Australia], 0.2 mM dNTPs (Roche Diag-

nostics, New Zealand), 1 U Platinum Taq DNA polymerase

(Invitrogen, Auckland, New Zealand), 4 mM MgCl2 (Invitro-

gen), 0.6 mg and nonacetylated bovine serum albumin (Sig-

ma, Auckland, New Zealand). Thermal cycling conditions

were: 94 1C for 30 s, followed by 55 1C for 45 s, 50 1C for 30 s,

72 1C for 2 min, repeated for 30 cycles with an initial

denaturation of 94 1C for 2 min, and a final extension of

72 1C for 7 min. PCR were run on an Eppendorf master-cycler.

PCR products were visualized on a 1.5% agarose gel, purified

using a High Pure PCR product purification kit (Roche

Diagnostics) and sequenced using the BigDye Terminator

v3.1 Cycle Sequencing Kit (Applied Biosystems). Sequencing

used primers 27F, 809R, 322 and 340. The sequences generated

during this work were deposited in the NCBI GenBank

database (refer to Table 1 for 16S rRNA gene accession

numbers) under accession numbers GU018020–GU018032

(ITS region). Amplification of the ITS was only undertaken on

isolates that fell within the Phormidium clade (based on 16S

rRNA gene sequences, Fig. 2).

To assess the microcystin-producing potential of each

culture, amplification of a region of the mcyE gene was

performed as described above using the HEPF and HEPR

Fig. 1. Locations of successfully isolated

cyanobacterial cultures.


97Benthic cyanobacteria in NZ

Tab

le1.

Origin

,cu

lture

num

ber

,m

orp

holo

gic

altr

aits

,to

xin

conte

nt

and

the

nea

rest

Gen

Ban

k(1

6S

rRN

Agen

ese

quen

ce)

iden

tifica

tion

mat

chfo

r36

ben

thic

cyanobac

terial

stra

ins

isola

ted

from

New

Zeal

and

rive

rsan

dla

kes

Culture

no.an

dlo

cation

Cel

l

wid

th

(mm

)

Cel

l

length

(mm

)

Dev

eloped

apic

alce

llC

ross

wal

ls

Anat

oxi

n

pro

duct

ion

(mg

kg�

1D

W)

Envi

ronm

enta

l,

ATX

/HTX

pro

duct

ion

(mg

kg�

1W

Wor

mg

kg�

1D

W)

Acc

essi

on

no.

IDof

nea

rest

mat

ch

(acc

essi

on

no.)

%ID

Sam

ple

loca

tion

Morp

hoty

pe

A

VU

W1,A

valo

nduck

pond

6.0

–7.8

2.4

–5.4

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451411

P.au

tum

nal

e;D

Q493873

98

E:2672100

N:5

999665

VU

W2,La

keH

enle

yO

utlet

6.6

–7.8

2.4

–3.6

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451409

P.au

tum

nal

e;D

Q493873

98

E:2735925

N:6

025095

VU

W3,W

ainuio

mat

aRiv

er4.8

–8.4

1.8

–3.6

Conic

al/c

apitat

e/

caly

ptr

a

––

GQ

451402

P.au

tum

nal

e;D

Q493873

98

E:2674410

N:5

990890

VU

W4,H

utt

Riv

er7.2

–7.8

4.2

–6.6

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

HTX

(4m

gkg�

1D

W)

dhA

TX(5

3m

gkg�

1D

W)

dhH

TX(9

mg

kg�

1D

W)

GQ

451410

P.au

tum

nal

e;D

Q493873

98

E:2674410

N:6

004090

VU

W5,W

aingongoro

Riv

er4.8

–6.0

2.4

–3.6

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

––

T.bourr

elly

i;A

B045897

98

E:2620350

P.au

tum

nal

e;EF

654081

98

N:6

199175

VU

W7,W

ainuio

mat

aRiv

er6.0

–6.6

3.0

–4.8

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451400

P.au

tum

nal

e;D

Q493873

98

E:2674410

N:5

990890

VU

W9,H

utt

Riv

er7.8

–9.6

3.6

–5.4

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451417

P.au

tum

nal

e:D

Q493873

98

E:2689075

N:6

010880

VU

W11,Pe

mbro

kero

ad6.0

–7.2

2.4

–4.8

Cap

itat

e/ca

lyptr

a–

NT

GQ

451408

P.au

tum

nal

e;D

Q493873

98

E:2657520

N:5

991385

VU

W14,H

utt

Riv

er7.2

–8.4

2.4

–3.6

Conic

al/c

alyp

tra

Gra

nula

r–

HTX

(9m

gkg�

1D

W)

dhA

TX(3

25

mg

kg�

1D

W)

dhH

TX(9

5m

gkg�

1D

W)

GQ

451399

P.au

tum

nal

e;D

Q493873

98

E:2670240

N:5

998870

VU

W16,Pe

loro

us

Riv

er6.6

–7.8

2.4

–4.2

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451398

P.au

tum

nal

e;D

Q493873

98

E:2558175

N:5

989825

VU

W17,M

angat

inoka

Stre

am6.0

–7.8

3.0

–4.2

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451407

P.au

tum

nal

e;D

Q493873

98

E:2752100

N:6

082600

VU

W18,M

akar

ewa

Riv

er6.0

–7.2

2.4

–3.6

Conic

al/c

apitat

e/

caly

ptr

a

––

GQ

451403

P.au

tum

nal

e;D

Q493873

98

E:2146875

N:5

420325

VU

W19,M

angar

oa

Riv

er6.0

–7.2

3.0

–4.8

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451416

P.au

tum

nal

e;D

Q493873

98

E:2688625

N:6

010315

VU

W20,Ran

gat

aiki

Riv

er6.6

–8.4

3.6

–4.8

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451415

P.au

tum

nal

e;D

Q493873

99

E:2835500

N:6

304300

VU

W21,W

hak

atan

eRiv

er7.2

–8.4

3.0

–5.4

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451432

P.au

tum

nal

e;D

Q493873

98

E:2860600

T.bourr

elly

i;A

B045897

98

N:6

312550

VU

W22,W

aim

ana

Riv

er7.2

–8.4

3.0

–6.0

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451406

P.au

tum

nal

e;D

Q493873

98

E:2869700

N:6

320350



VU

W23,G

odle

yRiv

erTr

ibuta

ry6.6

–8.4

3.0

–4.8

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451422

P.au

tum

nal

e;D

Q493873

98

E:2310125

T.bourr

elly

i;A

B045897

98

N:5

720475

VU

W24,Tu

kitu

kiRiv

er6.0

–9.6

3.0

–4.8

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

–G

Q451426

P.au

tum

nal

e;D

Q493873

98

E:2846550

T.bourr

elly

i;A

B045897

98

N:6

158400

P.au

tum

nal

e;EF

654081

98

CY

N47,A

shle

yRiv

er6.0

–7.2

3.0

–6.0

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

ATX

/HTX

GQ

451401

P.au

tum

nal

e;D

Q493873

98

E:2447450

N:5

774580

CY

N48,A

shle

yRiv

er6.0

–7.2

3.0

–6.0

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

ATX

/HTX

GQ

451404

P.au

tum

nal

e;D

Q493873

98

E:2475800

N:5

769700

CY

N49,H

utt

Riv

er5.4

–6.6

2.4

–4.2

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

ATX

/HTX

GQ

451405

P.au

tum

nal

e;D

Q493873

98

E:2670240

N:5

998870

CY

N52,Ran

gat

aiki

Riv

er4.2

–6.0

3.0

–5.4

Conic

al/c

apitat

e/

caly

ptr

a

ATX

(1000

mg

kg�

1)

ATX

(200

mg

kg�

1W

W)

GQ

451424

T.bourr

elly

i;A

B045897

99

E:2845850

N:6

350050

CY

N53,Ran

gat

aiki

Riv

er4.2

–6.0

3.0

–5.4

Conic

al/c

apitat

e/

caly

ptr

a

ATX

(1000

mg

kg�

1)

ATX

(200

mg

kg�

1W

W)

GQ

451413

T.bourr

elly

i;A

B045897

99

E:2845850

N:6

350050

CY

N55,Rodin

gRiv

er8.4

–9.6

5.4

–6.6

Conic

al/c

apitat

e/

caly

ptr

a

Gra

nula

r–

NT

GQ

451414

P.au

tum

nal

e;D

Q493873

99

E:2523465

N:5

977465

Morp

hoty

pe

B

VU

W8,A

kata

raw

aRiv

er9.6

–13.2

1.8

–4.2

Conic

al/c

apitat

e/

caly

ptr

a

–Tr

ace

leve

lsG

Q451420

P.au

tum

nal

e;D

Q493873

99

E:2686195

N:6

010975

VU

W10,W

ainuio

mat

aRiv

er8.4

–12

2.4

–4.8

Thic

ken/c

alyp

tra

Gra

nula

r–

NT

GQ

451419

P.au

tum

nal

e;D

Q493873

99

E:2678253

N:5

992345

VU

W12,W

ainuio

mat

aRiv

er9.6

–12

2.4

–4.2

Thic

ken/c

alyp

tra

Gra

nula

r–

–G

Q451418

P.au

tum

nal

e;D

Q493873

99

E:2678253

N:5

992345

Morp

hoty

pe

C

VU

W30,W

aika

toRiv

er9.0

–15.6

1.2

–4.2

Rounded

Gra

nula

r–

––

E:2776835

Slig

htly

rest

rict

edN

:6278400

Morp

hoty

pe

D

CY

N38,Red

Hill

sTa

rn3.6

–4.2

2.4

–4.2

Rounded

/conic

alSl

ightly

gra

nula

r–

NT

GQ

451428

P.m

urr

ayi;

DQ

493872

98

E:2513300

98

N:5

952770

CY

N39,Red

Hill

sTa

rn3.6

–4.2

2.4

–4.2

Rounded

/conic

alSl

ightly

gra

nula

r–

NT

GQ

451429

P.m

urr

ayi;

DQ

493872

98

E:2513300

N:5

952770

Morp

hoty

pe

E

VU

W13,M

angat

aurh

iriS

trea

m3.6

–4.8

3.6

–4.8

Rounded

––

GQ

451421

Sym

plo

casp

.;EU

249122

94

E:2701830

M.p

aludosu

s;EF

654090

93

N:6

454770

M.ch

thonopla

stes

;

EF654045

93

Morp

hoty

pe

F,G

,H

and

I

VU

W6,W

ainuio

mat

aRiv

er1.8

–2.4

1.2

–2.4

Rounded

Slig

htly

rest

rict

ed–

–G

Q451430

Pseu

dan

abae

na

trem

ula

;

AF2

18371

93

E:2674410

Lepto

lyngbya

frig

ida;

AY

493611

93

N:5

990890



primers (Jungblut & Neilan, 2005). PCR products were

visualized on a 1.5% agarose gel.

Sequence alignment and phylogeneticanalysis

The 16S rRNA gene sequences and ITS sequences were

aligned using CLUSTAL W in MEGA 4 (Tamura et al., 2007).

Pair-wise distances were calculated using the Jukes–

Cantor method and pairwise deletion used to account

for sequence length variation or gaps. Phylogenetic trees

were constructed using a neighbour-joining algorithm

(Saitou & Nei, 1987) and Tamura–Nei distance estimates.

Bootstrap analyses of 1000 iterations were performed to

identify the node support for the consensus trees.

Anatoxins analysis

Subsamples of all cyanobacterial strains were lyophilized

(FreeZone6, Labconco). Lyophilized material (100 mg)

was resuspended in 10 mL of double-distilled water

(DDW) containing 0.1% formic acid and sonicated (Cole

Parmer 8890, Biolab) for 15 min. Samples were centri-

fuged at 4000 g for 10 min. The procedure was repeated

using 5 mL DDW and the supernatants combined.

All cyanobacterial strains were analysed for ATX, HTX

and their degradation products using LC-MS. Anatoxins

were separated by LC (Acquity UPLC, Waters Corp., MA)

using a 500 1-mm Acquity BEH-C18 (1.7 mm) column

(Waters Corp.). Both the mobile phase A (water) and

mobile phase B (acetonitrile) contained 0.1% formic acid

and were used at a flow of 0.3 mL min�1, isocratic for

1 min at 100% A, followed by a rapid gradient from 100%

A to 50% A/50% B over 2 min. The injection volume was

5 mL. The Quattro Premier XE mass spectrometer

(Waters-Micromass, Manchester) was operated in the

ESI1 mode with capillary voltage 0.5 kV, desolvation gas

900 L h�1, 400 1C, cone gas 200 L h�1 and cone voltage

25 V. Quantitative analysis was performed by multiple-

reaction monitoring using MS-MS channels set up for

ATX (166.154 149.1; Rt 1.0 min), HTX (180.24 163.15;

Rt c. 1.9 min), dihydroanatoxin-a (168.14 56; Rt

0.9 min), dihydrohomoanatoxin-a (182.14 57; Rt

c. 1.9 min), epoxyanatoxin-a (182.14 98) and epoxyho-

moanatoxin (196.14 140; Rt c. 1.9 min). The instrument

was calibrated with dilutions in 0.1% formic acid of

authentic standards of ATX (A.G. Scientific, CA).

Results

Environmental samples and strain isolation

The majority of the 31 cyanobacterial mats sampled were

collected from black/green/brown leathery matsTab

le1.

Continued

.

Culture

no.an

dlo

cation

Cel

l

wid

th

(mm

)

Cel

l

length

(mm

)

Dev

eloped

apic

alce

llC

ross

wal

ls

Anat

oxi

n

pro

duct

ion

(mg

kg�

1D

W)

Envi

ronm

enta

l,

ATX

/HTX

pro

duct

ion

(mg

kg�

1W

Wor

mg

kg�

1D

W)

Acc

essi

on

no.

IDof

nea

rest

mat

ch

(acc

essi

on

no.)

%ID

Sam

ple

loca

tion

VU

W15,W

hak

atik

eiRiv

er1.8

–2.4

2.4

–4.2

Rounded

/conic

al–

Trac

ele

vels

GQ

451431

Osc

illat

oria

limnet

ic;

AJ0

07908.

99

E:2681885

Lim

noth

rix

redek

ei;

AJ5

80007

99

N:6

008350

Pseu

dan

abae

na;

AM

259269

99

VU

W26,W

hak

atan

eRiv

er1.2

1.2

–3.6

Cal

yptr

aC

onst

rict

edN

T–

GQ

451427

Pseu

dan

abae

na

sp.;

AM

259268

97

E:2860600

Art

hro

nem

agyg

axia

na;

AF2

18370

97

N:6

312550

VU

W28,H

utt

Riv

er1.2

–1.8

1.2

–3.6

Rounded

/conic

alSl

ight

const

rict

ion

NT

NT

GQ

451412

Pseu

dan

abae

na

sp.;

AM

259268

97

E:2670240

N:5

998870

Morp

hoty

pe

J

VU

W31,W

aika

toRiv

er3.0

–3.6

3.6

–4.2

Res

tric

ted

––

GQ

451425

Nost

oc

musc

oru

m;

AM

711524

99

E:2776835

N:6

278400

P.,Ph

orm

idiu

m;T.

,Ty

chonem

a;M

.,M

icro

cole

us;

1/�

,det

ecte

d/n

ot

det

ecte

d;N

T,not

test

ed;dhH

TX,dih

ydro

hom

oan

atoxi

n.



consistent with that described for Phormidium (Biggs &

Kilroy, 2000; Wood et al., 2007). Only two other mat types

were found and collected: green gelatinous Nostoc colonies

and one brittle brown filamentous mat. Preliminary micro-

scopic observation of the leathery mats confirmed that they

were comprised almost entirely of filamentous Phormidium

sp.. Representatives of Oscillatoria were observed among two

of the Phormidium-dominated mats. Additionally, Pseuda-

nabaenaceae were observed in at least 10 of the samples, but

these species proved difficult to isolate and culture. The

brittle brown mat was a proliferation of two species from the

Pseudanabaenaceae. Of the Nostoc colonies observed, only

one could be cultured and described. This strain of Nostoc

was found to be growing with an Oscillatoria sp.; together,

they were the only species collected from a geothermal

location. Thirty-six unicellular cyanobacterial strains were

successfully isolated and on-grown. Within all mat samples,

a mixture of diatoms was also observed. Melosira, Cymbella,

Frustulia and Gomphonema were the most prominent.

Microscopic characterization of isolates

Morphological characteristics including cell dimensions,

apical cell profile, cross wall configuration and reproductive

CYN47 Ashley River

VUW17 Mangatinoka River

CYN49 Hutt River

VUW21 Whakatane River

VUW16 Pelorous River

CYN48 Ashley River

VUW18 Makarewa River

VUW24 Tukituki River

VUW22 Waimana River

VUW23 Godley River

VUW14 Hutt River

VUW7 Wainuiomata River


VUW5 Waingongoro River

VUW4 Hutt River

VUW2 Lake Henly

VUW1 Avalon Duck Pond

VUW11 Pembroke Road Stream

CYN53 Rangataiki River

CYN52 Rangataki River

Tychonema bourrellyi AB045897

Phormidium autumnale EF654081

Microcoleus antarcticus AF218373

Phormidium autumnale DQ493874

Phormidium autumnale DQ493873

CYN55 Roding River

VUW20 Rangataiki River

VUW19 Mangaroa River

VUW9 Hutt River



VUW8 Akatarawa River

Morphotype B68

68

57

66

89

8569

69

75

57

69

55

Morphotype A

Fig. 2. Phylogenetic tree of the Phormidium autumnale group based on the 16S rRNA gene sequences (647 bp) and obtained using the neighbour-

joining method. Bootstrap values 4 50% are noted at the nodes. The different morphotypes in this study are in bold. The remainder of the tree is

shown in Fig. 5.



structures for each isolate are given in Table 1. All isolates

were found to have traits in common with Oscillatoriales,

with the exception of VUW31, which was placed in the

Nostocales.

Cells from the majority of isolates were characterized by

being isodiametric or slightly shorter than wide. Cells were

generally 4–8.5 mm wide. Trichomes were motile and

straight with well-defined apical cells with a calyptra (thick-

ened membrane) (Fig. 3a–h). Those isolates sharing these

morphological characteristics were identified as P. autum-

nale [(Agardh) Trevisan ex Gomont 1892] (Komarek &

Anagnostidis, 2005; McGregor, 2007; Wood et al., 2007)

and assigned to Morphotype A (Table 1, Fig. 31–h). Twenty-

four strains VUW1–5, 7, 9, 11, 14, 16–24, CYN47–49,

CYN52–53 and 55 were included in this designation.

Morphological variation was observed among strains in

apical cell morphology, cell-wall granulation, sheath pre-

sence or absence and thickness.

Stains VUW8, 10 and 12 were morphologically similar to

strains from Morphotype A; however, the cell structure was

discoid (width, 8.4–13.2 mm; length, 1.8–4.8 mm) and each

had a distinctive, rounded to hemispherical calyptra (Fig. 3i).

Isolates were identified as belonging to the genus

Oscillatoria possessing a distinctly thickened apical cell with

calyptra, no constrictions at cross walls and incomplete

cross-wall formation during cell division (the main generic

feature) (Komarek & Anagnostidis, 2005). These three

strains were assigned to Morphotype B. The only other

strain identified as Oscillatoria was VUW30. This strain

(Morphotype C) was distinctively granular, generally

straight, possessing a discoid cell structure (width,

8.4–15.6 mm; length, 2.2–3.8 mm) and was slightly con-

stricted at cross walls (Fig. 4a).

Morphotype D (Fig. 4b) was represented by strains CYN38

and 39, which were assigned to Phormidium murrayi (West

and West 1911) (Anagnostidis & Komarek, 1988; Comte et al.,

2007), with narrow, isodiametric cells (width, 3.6–4.2mm;

length, 2.4–4.2mm), a prominent sheath (not always present)

and conical/rounded apical cells with no calyptra.

Strain VUW13 (Morphotype E) unfortunately stopped

growing in culture before a full taxonomic classification

could be provided; however, preliminary identification

assigned this to the Phormidiaceae. It was distinctly different

from all the other morphotypes.

(a)

(f)

(e)

(d)

(c)

(b)

(i)

(g)

(h)

Fig. 3. Photomicrographs of nine different Phormidium autumnale strains, (a–h) Morphotype A and (i) Morphotype B. Photomicrographs images are all

bright field, with the exception of (c) and (d), which were taken using Nomarski optics. (a) VUW18 Makarewa River, (b) VUW9 Hutt River, (c) VUW19

Mangaroa River, (d) CYN55 Roding River, (e) CYN53 Rangataiki River, (f) VUW1 Avalon Duck Pond, (g) VUW17 Mangatinoka Stream (Lugol’s preserved),

(h) VUW24 Tukituki River and (i) VUW10 Wainuiomata River (red border, only representative from Morphotype B). All are described in Table 1. Scale

bars = 20mm.



Single strains from the family Pseudanabaenaceae were

assigned as Morphotypes F, G, H and I, and all differed

slightly in morphology. Strains VUW6 (Morphotype F,

Fig. 4c) and VUW28 (Morphotype I) were assigned to

genus Leptolyngbya. Morphotype F was found in tightly

tangled mats. Trichomes were slightly constricted at cross

walls. Cells were isodiametric (width, 1.8–2.4 mm; length,

1.2–2.4 mm). Apical cells were rounded/conical and sheath

was absent. Strain VUW28 (Morphotype I) formed a

dense tangled mat, possessing no aerotopes, having a slight

constriction, sheath sometimes present and a conical apical

cell. Strain VUW15 (Morphotype H) was the only species

isolated from the brittle brown mat. The strain was found to

form benthic mats, comprising aerotopes at the septa, with a

slight constriction at cross walls and cells two to three times

longer than wide. Strain VUW26 (Mophotype H) was found

to be from the Pseudanabaena genus, defined by very brittle

filaments and cell wall constriction. It formed a very loose

‘soup’ of filaments.

Finally, strain VUW31 was the only representative of

Nostoc. Possessing distinctive akinetes and heterocytes, this

strain was identified as Nostoc muscorum and designated as

Morphotype J (Fig. 4d).

Genotypic analyses and comparison withmorphological designations

The 16S rRNA gene sequences (647 bp) for all 36 cultures,

with the exception of culture VUW30, were used to con-

struct a neighbour-joining tree to determine the phyloge-

netic relationship between isolates and other cyanobacterial

16S rRNA gene sequences obtained from GenBank (Figs 2

and 5).

Twenty-seven of the cultured isolates clustered together

with 100% bootstrap support (Fig. 2). Using BLASTN, all 27

sequences in this group matched at 4 98% sequence

homology with P. autumnale representatives from GenBank

(all GenBank representatives were from peer-reviewed pub-

lications). The 27 isolates, however, shared a sequence

similarity o 97%. They consisted of all strains from Mor-

photypes A and B. Within this clade, there were a number of

slight genotype variations, resulting in the formation of a

number of subclades, and these aligned with the slight

morphological differences observed in this group (Fig. 2).

Interestingly, strains from Morphotype B were found in a

larger clade including strains from Morphotype A.

Strains CYN38 and 39 (Morphotype D), which shared

identical sequences, clustered together in a subgroup with P.

murrayi (DQ493872 and AY493627) and Microcoleus glaciei

(formerly P. murrayi, AF218374) with 99% bootstrap sup-

port and 4 97% sequence similarity (Table 1; Fig. 5). Other

strains on the sister branch were distant, sharing o 93%

similarity.

Strain VUW13 (Morphotype E) clustered most closely

with Microcoleus, Symploca and Lyngbya sp.; however, this

was supported with only 64% bootstrap and o 95%

sequence similarity (Table 1; Fig. 5). The nearest Phormidium

strains (P. murrayi) exhibited o 93% sequence similarity.

With the exception of strain VUW6, strains of the

Pseudanabaenaceae could not be further defined by phylo-

genetic analysis (Fig. 5). Strain VUW6 (Morphotype F)

was found to cluster with Leptolyngbya frigida sequences

(AY493611) and Pseudanabaena tremula (AF218371).

(a) (c)

(d)(b)

Fig. 4. Photomicrographs of four strains from

contrasting genera. (a) VUW30 Waikato River

(Morphotype C), (b) CYN38 Red Hills Tarn

(Morphotype D), (c) VUW6 Wainuiomata River

(Morphotype G) and (d) VUW31 Waikato

River (Morphotype J). Scale bars = 20 mm.

All images are bright field.



Phormidium autumnale group (See Fig. 2)Phormidium uncinatum EF654086

Planktothrix sp. GQ451423.

Planktothrix rubescens AB045925

Planktothrix rubescens AJ132252

Planktothrix agardhii AB045923

Oscillatoria sancta AF132933

Limnothrix redekei AJ505942

Oscillatoria duplisecta AM398647

Oscillatoria princeps AB045961

Phormidium tergestinum EF654083

Phormidium sp. DQ235808




CYN38 Red Hills Tarn Morphotype D

CYN39 Red Hills Tarn Morphotype D

Phormidium murrayi DQ493872

Phormidium murrayi AY493627

Microcoleus glaciei AF218374

Microcoleus chthonoplastes EF654059

Microcoleus chthonoplastes EF654060

VUW13 Man gataurhiri River Morphotype ESymploca sp. EU249122

Lyngbya bouillonii FJ147302

Lyngbya majuscula FJ356670

Microcoleus paludosus EF654090

Cylindrospermopsis raciborskii AF516746

Cylindrospermum sp. AJ133163

Anabaena lemmermannii AJ293113

VUW31 Waikato River Nostoc muscorum AM711524

Pseudanabaena tremula AF218371

Leptolyngbya frigida AY493611


Morphotype J

Morphotype FLimnothrix redekei AJ505941



VUW26 Whakatane River Morphotype H

VUW28 Hutt River Morphotype IArthronema gygaxiana AF218370

VUW15 Whakatikei River Morphotype GLimnothrix redekei AB045929


Oscillatoria limnetica AJ007908

Pseudanabaena sp. AM259269

Escherichia coli CU928160

60

96

54

100

98100

79100

100

8383

100

100

100

100

99

100

100

64

92

98

50

100

100

64

87

85

61

64

50

100

Fig. 5. Phylogenetic tree based on the 16S rRNA gene sequences (647 bp) and obtained using the neighbour-joining method. Bootstrap values 4 50%

are noted at the nodes. The different morphotypes from this study are in bold. The Phormidium autumnale section of the tree is shown in Fig. 2.



The remaining three strains (VUW15, 26 and 28; Morpho-

types G, H and I) clustered together with GenBank se-

quences Arthronema gygaxiana (AF218370), Oscillatoria

limnetica (AJ007908), Pseudanabaena sp. (AM259269) and

Limnothrix redekei (AB045929 and AJ580007) with 99%

bootstrap support and 4 98% sequence similarity.

Finally, strain VUW31 (Morphotype J) was found to

cluster with other heterocytous species and had 99% node

support with Nostoc moscorum (AM711524).

ITS sequences (727 bp) were successfully obtained for 12

isolates from within the Phormidium clade (VUW 2, 4, 8, 9,

16–18, 24, CYN 47–49 and 55). These sequences were used

to construct a neighbour-joining tree. In general, the

phylogenetic relationships observed in this tree matched

those observed in the phylogenetic analysis of 16S rRNA

gene sequences (data not shown). The ITS sequences did not

provide any additional information to allow finer scale resolu-

tion of intraspecies identification within the P. autumnale

clade.

Cyanotoxin analysis

LC-MS analysis identified anatoxins in seven of the 31

environmental samples (Table 1). All seven mats were

dominated by Phormidium. Interestingly, ATX was detected

in only two of the 36 strains (CYN52 and CYN53), while

HTX and its degradation products were not detected. Both

strains produced ATX at 1000 mg kg�1 freeze-dried weight.

Both strains CYN52 and CYN53 were isolated from the

same environmental sample sourced from the Rangataiki

River and identified by morphology and phylogenetic

analysis as P. autumnale. These two strains also formed their

own clade in the phylogenetic tree sharing 100% sequence

similarity (Fig. 2).

No isolates were found to contain the mcyE gene, with

only the positive control testing positive.

All strains isolated in this study have been cryopreserved

successfully using the methods of Wood et al. (2008), and

these are banked and maintained in the Cawthron Institute

culture collection (http://www.cawthron.org.nz/seafood-sa

fety-biotechnology/micro-algae-culture-collection.html).

Discussion

Morphological investigation revealed the presence of nine

different morphotypes, all from the Oscillatoriales and one

(Morphotype J) from the Nostocales.

Morphotypes A and B

Phormidium autumnale is known to have a cosmopolitan

distribution and has been identified in many different

habitats (Komarek & Anagnostidis, 2005; Palinska & Mar-

quardt, 2008). It has unusually broad morphological and

physiological characteristics (Palinska & Marquardt, 2008).

These features were also observed in this study. Morphotype

A consisted of 24 isolates that were identified as P. autum-

nale by polyphasic assessment. These 24 isolates could not

be further separated based on morphological criteria. Con-

siderable morphological variation was, however, observed

between isolates, but this variation was confined within the

broad P. autumnale definition (Komarek & Anagnostidis,

2005). Cell granulation, apical cell profile and sheath

presence were all found to vary. Sheath production, tradi-

tionally used for the systematic classification of Oscillator-

ialean (Anagnostidis & Komarek, 1988; Komarek &

Anagnostidis, 2005; Palinska & Marquardt, 2008), has been

shown to be subject to the direct effects of both environ-

mental and culture conditions (Rippka et al., 1979; Whitton,

1992). Apical cell structures (in particular, the calyptra) vary

in trichomes of different ages and are rarely seen in culture

(Komarek & Anagnostidis, 2005). The observed variation

seen in this study is in contrast to Palinska & Marquardt

(2008), who demonstrated that 10 P. autumnale ssp. exhib-

ited a relatively similar morphology in culture.

Intraspecific identification was therefore not possible in

this group by morphology. However, the morphological

heterogeneity observed in Morphotype A was represented

by a number of genotype variations in the phylogenetic

analysis. These genotype variations resulted in the formation

of 10 different clades, indicating the presence of subspecies

within this morphotype. This is consistent with Comte et al.

(2007), who were unable to separate their P. autumnale

strains Arct-Ph5 and Ant-Ph68 based on morphological

characteristics, but identified two subspecies by genetic

analysis (16S rRNA gene sequence). Strains CYN52 and 53

formed their own clade within Morphotype A and were the

only anatoxin producers in this lineage. Isolation of further

anatoxin-producing strains is required to determine whether

this divergence is consistent for all anatoxin producers.

Unexpectedly, the three strains of Morphotype B, identi-

fied as Oscillatoria by morphology, formed their own clade

within this P. autumnale lineage. Morphotype B has disc-like

cells and a hemispherical calyptra. These features contrast

with those of Morphotype A. Palinska & Marquardt (2008)

found that cell size varies not only between the strains but

also within the strains of P. autumnale. This was also found

in the current study, where Morphotype B possessed cell

widths ranging between 8.4 and 13.2mm. This is larger and

with a greater variation than that described previously for

this species and differs markedly from Morphotype A.

Apical cell structure (calyptra) furthermore has been recog-

nized as a stable morphological structure used for distin-

guishing species (Komarek & Anagnostidis, 2005). In this

study, two distinct calyptra types were observed among

Morphotypes A and B (Fig. 3), indicating that these two

morphotypes should be maintained as two distinct species.



http://www.cawthron.org.nz/seafood-safety-biotechnology/micro-algae-culture-collection.html

http://www.cawthron.org.nz/seafood-safety-biotechnology/micro-algae-culture-collection.html

The phylogenetic analysis (16S rRNA gene sequences)

showed that the P. autumnale lineage consisted of all isolates

from Morphotypes A and B. Additionally, GenBank repre-

sentatives Microcoleus sp. and Tychonema bourrellyi were

included in this clade (Fig. 2). This clade was supported

by 99% bootstrap support; however, together, they shared

o 97% sequence homology. Previous studies have used

sequence homologies of 4 97.5% as a threshold for bacter-

ial species definition, while 95% has been used as a genus

barrier for the 16S rRNA gene (Stackebrandt & Goebel,

1994; Casamatta et al., 2005; Palinska & Marquardt, 2008).

This standard led Palinska & Marquardt (2008) to conclude

that their two morphologically similar P. autumnale groups

that shared o 97% sequence homology may be two differ-

ent species of the same genus. Thus, the 10 different clades

observed in this study may represent a number of different

species of the same genus.

GenBank sequences for Microcoleus antarcticus and

T. bourrellyi also clustered in the P. autumnale lineage. This

is consistent with previous research (Willame et al., 2006;

Comte et al., 2007; Palinska & Marquardt, 2008). Palinska &

Marquardt (2008) demonstrated the strong morphological

similarities between these genera, and concluded that the

phenotypic criteria are too uncertain and overlapping for a

clear distinction between P. autumnale, T. bourrellyi and

Microcoleus sp..

Morphotype D

Both strains CYN38 and 39 were identified as P. murrayi by

morphologic and phylogenetic analyses. This morphotype

comprises long narrow trichomes, isodiametric cells and a

prominent sheath that separates it from Morphotypes A and

B. In the phylogenetic analysis, the two strains were found to

cluster in a subgroup supported by 99% bootstrap support

with three Antarctic P. murrayi strains (DQ493872 and

AY493627 and Microcoleus glacei formerly P. murrayi,

AF218374). To our knowledge, this is the first recorded case

of P. murrayi occurring outside of Antarctica and dispels

previous views that this morphotype is endemic to Antarc-

tica (Casamatta et al., 2005). Furthermore, it raises interest-

ing questions about the dispersal, habitat and distribution of

this species.

Recently, there has been conjecture over the classification

of the P. murrayi, which may cluster in the Microcoleus genus

and not Phormidium (Casamatta et al., 2005; Comte et al.,

2007). The sister branch to the P. murrayi clade in this

study’s tree comprises species from Microcoleus and Lyng-

bya, but none from Phormidium. GenBank sequence homo-

logies for morphotype D were found to align more closely

with Microcoleus sp. than Phormidium sp.. These results are

consistent with previous investigations (Casamatta et al.,

2005; Taton et al., 2006; Comte et al., 2007) that show

P. murrayi aligning in clades other than Phormidium.

Casamatta et al. (2005) reassigned P. murrayi to M. glaciei

on the basis of their findings. In accordance with Comte

et al. (2007), the genetic coincidence of our strains with

those of the genus Microcoleus needs further clarification, a

result that has now been demonstrated in three separate

studies.

Morphotypes C, F, G, H, I and J

Strain VUW30 (Morphotype C) was identified as

Oscillatoria sp. based solely on morphological characteriza-

tion. Strain VUW13 (Morphotype F) identified by mor-

phology as Phormidium aligns with non-Phormidium

genera (Lyngbya, Symploca, Microcoleus) by phylogenetic

analysis. Casamatta et al. (2005) suggest that Phormidium

sp. aligning in this non-Phormidium group are most likely

to be from a different genus. Morphotypes F, G, H and I

were identified as members of the Pseudanabaenaceae.

Komarek & Anagnostidis (2005) state that clear separation

of genera from this family by molecular analyses is not well

defined. This is consistent with the results found in this

study, with a number of different genera found in the same

clade. Further morphological classification using a greater

phenotypic criterion is needed in the analysis of these

morphotypes.

Cyanotoxin production

In routine testing of Phormidium mats around New Zeal-

and, it has been found that the occurrence, concentration

and variants of anatoxins are unpredictable (S.A. Wood,

unpublished data). In a study of anatoxin-producing

benthic cyanobacteria in the Tarn River, France, Cadel-Six

et al. (2007) found that Phormidium strain Fil.2Da FY

produced ATX even though it was isolated from an appar-

ently nontoxic sample. They concluded that this result may

be due to both toxic and nontoxic representatives of the

same phenotype occurring at a single site, and fortuitously,

they were able to isolate a filament that proved to be a toxin

producer. In contrast, in this study, we isolated a number of

nontoxic strains from environmental samples that were

known to contain anatoxins (Table 1). Anatoxin was only

detected in two of our strains (CYN52 and 53), which were

both isolated from a cyanobacterial mat that had been

collected in response to a dog neurotoxicosis on the Ranga-

taiki River.

Our P. autumnale isolates showed a large radiation of

subspecies, and it seems likely that mats are comprised of a

mixture of strains, only some of which have the ability to

produce anatoxins. Therefore, the likelihood of obtaining an

anatoxin-producing strain is dependent on the filament

isolated and its relative abundance in the mat. This is

consistent with patterns observed in planktonic



cyanobacteria, for example, Vezie et al. (1998), showed that

blooms of planktonic microcystin-producing cyanobacteria

are comprised of toxic and nontoxic strains. The hypothesis

that mats are comprised of toxic and nontoxic genotypes

may also explain the significant variation in the anatoxin

concentrations observed in environmental samples from

around New Zealand, i.e. samples with high concentrations

of anatoxins are likely to be dominated by toxic genotypes,

while those with low concentrations may contain only a few

filaments of toxic genotypes. Different anatoxin variants are

also likely to be indicative of the presence of different toxin-

producing genotypes. In this study, HTX was not produced

by any of the isolates, even though it was detected in the

initial screening of environmental samples (Table 1). It is

also possible that some of the isolates in the present study

did not produce detectable anatoxins under laboratory

conditions; however, this hypothesis seems unlikely, given

the favourable laboratory conditions for growth.

The lack of microcystin producers within isolates in this

study was surprising, given that low concentrations of

microcystins have been detected previously (via ADDA-

ELISA) in benthic cyanobacterial mats in New Zealand

(Hamill, 2001; Wood et al., 2006). Phormidium sp. elsewhere

have also been shown to produce microcystins (e.g. Mo-

hamed et al., 2006; Izaguirre et al., 2007). Further isolation

of strains from environmental samples known to contain

microcystins may help in the identification of benthic

microcystin species in New Zealand.

Conclusions

The results of this study revealed that P. autumnale is the

predominant cyanobacterium in benthic mat proliferations

in New Zealand rivers. Polyphasic analysis showed that these

mats are comprised of a number of different P. autumnale

spp. and that only certain subspecies produce anatoxins. The

ATX-producing P. autumnale strain identified in this study

was unique at the 16S rRNA gene sequence level. Isolation of

further toxic strains would be required to assess whether

anatoxin production is limited to only this strain across New

Zealand and to investigate whether this difference could be

used in the development of a molecular-based diagnostic

tool. The marked morphological variation observed within

P. autumnale highlights the difficulties in identifying cyano-

bacteria from the Oscillatoriaceae to the species level based

solely on morphology. The use of a polyphasic approach is

recommended. However, the number of 16S rRNA gene

sequences currently deposited in worldwide databases is

limited and larger scale systematic projects will be required

to fully resolve current taxonomic uncertainties. This study

emphasizes the paucity of information currently available

on the diversity and toxin capabilities of New Zealand

benthic cyanobacteria.

Acknowledgements

This research was funded by the Greater Wellington Regio-

nal Council science research fund, the Victoria University

of Wellington strategic research fund, the New Zealand

Foundation for Research Science and Technology (FRST)

(CAWX0703), an Andy Ritchie Memorial Trust scholarship to

M.W.H. and a FRST postdoctoral fellowship (CAWX0501) to

S.A.W. The authors thank Michael Boundy and Roel Van

Ginkel (Cawthron Institute) for technical assistance with

anatoxin analysis.

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Polyphasic assessment of fresh-water benthic mat-forming cyanobacteria isolated from New Zealand

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