Coral Reefs (2005) 24: 57–66
DOI 10.1007/s00338-004-0442-6
R EP O RT
T. L. Shearer Æ C. Gutiérrez-Rodrı́guez Æ M. A. Coffroth
Generating molecular markers from zooxanthellate cnidarians
Received: 28 April 2003 / Accepted: 4 May 2004 / Published online: 5 November 2004
Springer-Verlag 2004
Abstract Genetic techniques are providing tools that are
necessary to answer questions concerning the ecology
and evolution of cnidarians that, until recently, could
not be easily addressed. In developing molecular markers for cnidarians with algal symbionts (zooxanthellae),
however, caution must be used to ensure the markers in
question are derived from the cnidarian host and not
zooxanthellae. Unless the DNA template is from
asymbiotic tissue, both host and symbiont genomes are
present in the DNA template and zooxanthella-specific
markers are often inadvertently generated. Steps should
be taken to minimize the contamination by zooxanthella
DNA in the template, and the origin of the molecular
marker (host or symbiont) must be verified. Including
zooxanthella-specific markers in analyses for cnidarians
will confound interpretations of the results as biogeographic and phylogeographic patterns of zooxanthellae
do not necessarily reflect those of the host.
Keywords Cnidaria Æ Zooxanthellae Æ Molecular
markers Æ DNA sequencing Æ Microsatellite
Introduction
As the fate of coral reefs is becoming a concern,
molecular techniques are increasingly being used to answer ecological and evolutionary questions with regard
to corals and other cnidarians. Genetic markers have
Communicated by Ecological Editor P.F. Sale
T. L. Shearer (&) Æ C. Gutiérrez-Rodrı́guez Æ M. A. Coffroth
Department of Biological Sciences, University at Buffalo,
109 Cooke Hall, Buffalo, NY 14260, USA
E-mail: [email protected]
Tel.: +1-716-6452363
Fax: +1-716-6452975
C. Gutiérrez-Rodrı́guez
Department of Biological Sciences, Ohio University,
Athens, OH 45701, USA
been used to delineate population structure and gene
flow in cnidarian populations (e.g. Beiring 1997;
Coffroth and Lasker 1998; Gutiérrez-Rodrı́guez 2003;
Kim et al. 2004; Le Goff-Vitry et al. 2004; GutiérrezRodrı́guez and Lasker 2004a; MacKenzie et al. 2004; I.
Baums, unpublished data; E. Severance, unpublished
data; T. Shearer, unpublished data), provide insight into
evolutionary relationships among cnidarian species (e.g.
Bridge et al. 1992; France et al. 1996; Romano and
Palumbi 1996, 1997; Lopez et al. 1999; Medina et al.
1999; Chen and Yu 2000; Romano and Cairns 2000;
Fukami et al. 2000; Berntson et al. 2001; Chen et al.
2002), and determine how processes such as hybridization have influenced the evolution of extant cnidarians
(e.g. Hatta et al. 1999; van Oppen et al. 2000; Diekmann
et al. 2001; Vollmer and Palumbi 2002). However, the
close association of many cnidarians with symbiotic algae, intracellular dinoflagellates termed zooxanthellae,
can complicate molecular analyses of cnidarian species.
Since zooxanthellae reside within host cells, extraction
of DNA from adult cnidarian tissue yields a DNA
template containing both cnidarian and algal genomes.
Thus, investigating micro- and macroevolutionary processes in symbiotic cnidarians requires verification that
the genetic markers used are derived from the host and
not from zooxanthalla contamination. Depending on
the molecular technique, either host-specific primers
must be designed or rigorous testing must be conducted
to confirm that the markers are derived from the host.
Without rigorous testing to confirm that molecular
markers generated from adult cnidarian tissue are cnidarian in origin, caution should be taken in interpreting
the data. Unknowingly including zooxanthella markers
in analyses for cnidarians can mislead and confuse the
interpretations of the results. The primary intention of
this manuscript is to illustrate that molecular markers can
easily, frequently, and inadvertently be generated from
symbionts inhabiting cnidarians. Indeed, the potential for
this contamination has been recognized by many cnidarian biologists, since the most readily available specimen samples contain zooxanthellae. Thus, the secondary
58
intention is to describe our use of various methodologies
available in the literature that have been developed to
minimize this concern and provide guidelines to develop
molecular markers for zooxanthellate cnidarians.
Specificity of molecular marker primers
Molecular markers are broadly categorized into two
groups: those that utilize primers specific to a particular
gene or species (i.e. nucleotide sequencing, microsatellites)
and those with arbitrary primers used in fragment analyses (i.e. randomly amplified polymorphic DNA (RAPDs), arbitrary fragment length polymorphisms (AFLPs),
inter-simple sequence repeats (ISSRs)). The strategy used
to control for the presence of zooxanthellae is not only
dependent on which type of marker will be utilized, but
also on the tissue available to generate the cnidarian
markers (i.e. symbiotic or asymbiotic). For molecular
markers in which host-specific primers can be designed
(i.e. sequencing and microsatellites), use of asymbiotic
gametes or larvae is ideal to eliminate the potential of
generating non-host markers and to avoid the rigorous
testing that is necessary to confirm host specificity of the
primers (as in Odorico and Miller 1997; Takabayashi et al.
1998; Hatta et al. 1999; Lopez et al. 1999; Medina et al.
1999; Chen and Yu 2000; Fukami et al. 2000; van Oppen
et al. 2000). Alternatively, design of cnidarian-specific
primers from sequences of close asymbiotic relatives (i.e.
Metridium senile, Tubastraea coccinea) ensures that the
appropriate taxon is amplified during the polymerase
chain reaction (PCR; as in Bridge et al. 1992; Chen and Yu
2000). When cnidarian-specific primers are unavailable,
sequences generated using ‘‘universal’’ primers can be
used to generate species- or cnidarian-specific primers
once these sequences have been properly verified as cnidarian in origin (see below; France et al. 1996; Chen et al.
2000; Berntson et al. 2001; Geller and Watson 2001). Once
cnidarian-specific primers are designed from zooxanthella-free tissue or from verified cnidarian sequences,
PCR amplification targets the cnidarian genome when the
DNA template contains both cnidarian and algal genomes.
The use of asymbiotic tissue is ideal for molecular
markers that utilize arbitrary primers (i.e. RAPDs,
AFLPs, ISSRs), in order to eliminate the potential of
including zooxanthella-derived bands in the analysis.
This, however, is usually impossible for population genetic studies, as 30–50 samples are a minimum statistical
requirement. It is, therefore, necessary to verify that the
resultant amplification products are cnidarian in origin
and not from contaminating zooxanthella DNA (see
below).
Minimizing zooxanthella contamination
Zooxanthella-free gametes are the preferred starting
material for generating cnidarian molecular markers,
however, obtaining gametes can be challenging for many
species. Obtaining zooxanthella-free larval tissue is not
possible for species in which zooxanthellae are transferred directly from the maternal colony to larvae (e.g.
Rinkevich and Loya 1979; Chornesky and Peters 1987;
Harii et al. 2002). Furthermore, adult samples are often
more readily available for DNA extraction. In situations
where symbiont-free tissue is unavailable to design hostspecific primers or for analyses involving arbitrary
primers, it is prudent to minimize cross-contamination
with zooxanthella DNA.
Using methods to enrich the DNA template toward a
higher proportion of host DNA relative to algal DNA
can increase the probability of amplifying host-specific
products during PCR. A potential problem with this
strategy is that the genome size of zooxanthellae may be
much larger than that of cnidarians [dinoflagellates: 2.2–
200 pg DNA per haploid nucleus (Spector 1984);
cnidarians: 0.23–1.85 pg DNA per haploid nucleus,
(Gregory 2001)]. Thus, the vast majority of zooxanthella
cells must be removed from the host tissue for the DNA
template to be host-biased. Although use of these techniques cannot guarantee generation of cnidarian markers exclusively, a higher proportion of these markers
may be of cnidarian origin as verified through various
methods (see below).
Centrifugation at low speed has routinely been used
to separate zooxanthellae for host extracts (e.g. Rowan
and Powers 1991; Lopez et al. 1999; LaJeunesse 2002).
In addition, separating intact zooxanthella cells from
host tissue by differential centrifugation using Percoll
gradients can also reduce zooxanthella contamination of
the host fraction (e.g. Tytler and Davies 1983; Stochaj
and Grossman 1997; Garson et al. 1998). However,
DNA released from any algal cells broken during the
early steps will contaminate the host DNA template.
In some cnidarians, such as sea anemones or some
soft corals, the distribution of zooxanthellae is heterogeneous throughout the host with few zooxanthellae
residing in areas such as the pedal disc or below the
surface layers (Fautin and Smith 1997; ten Lohuis et al.
1990). Use of tissue from these areas minimizes the
amount of algal contamination of the cnidarian DNA
template (ten Lohuis et al. 1990; Fautin and Smith 1997;
Pinto et al. 2000). Avoiding contamination by physically
removing algal cells from coral tissue under a microscope has yielded sufficient amounts of symbiont-free
tissue to design coral microsatellite primers (Maier et al.
2001).
Although the use of bleached coral tissue has not
been reported in the development of coral-specific
markers, by removing zooxanthella cells from the host
tissue through various bleaching methods (e.g. Vandermeulen et al. 1972; Steen and Muscatine 1987; Kinzie
et al. 2001; Belda-Baillie et al. 2002), the proportion of
zooxanthella DNA relative to coral DNA in the
extraction can be reduced. This may not be a preferred
method of obtaining uncontaminated cnidarian tissue as
bleaching is often impractical depending on the species,
59
and does not eliminate all algal cells from the host (Steen
and Muscatine 1987; Berner et al. 1993).
Verification of source of marker
For each putative cnidarian marker developed, it is
essential to confirm the origin of the amplification
products by testing primers against zooxanthella controls. Confirming the origin of the marker is especially
important when using ‘‘universal’’ primers, in designing
cnidarian-specific primers from symbiotic tissue (e.g.
sequencing and microsatellites) and in the various fragment analyses that utilize arbitrary primers (e.g. RAPDs, AFLPs, ISSRs), as DNA from both organisms can
be amplified during PCR, even with cnidarian-biased
templates (see results). When generating sequence data,
one approach to verify the source of the amplified
sequence is to determine phylogenetic affinity with
sequences from asymbiotic cnidarian relatives or other
animal taxa using database searches such as BLAST in
GenBank (Altschul et al. 1997). This technique has been
used to support the cnidarian origin of various cnidarian
sequences (e.g. Hunter et al. 1997; Snell et al. 1998; Chen
and Yu 2000; Chen et al. 2000; Berntson et al. 2001;
Diekmann et al. 2001). To determine which bands generated from adult tissue are attributed to zooxanthellae
in microsatellite and fragment analyses, comparisons
with banding patterns generated from zooxanthella-free
tissue and/or from pure zooxanthella cells can be made.
Due to intraspecific polymorphisms, banding patterns of
several host samples must be compared to several zooxanthella ‘‘control’’ samples to account for most, if not
all, of the observed bands. PCR products can be identical in size, but not homologous (e.g. Lynch 1988),
therefore, bands shared between host and zooxanthellae
should be cloned and sequenced to determine if
homologous regions are amplified or if zooxanthella
DNA is amplified from the host tissue. A second, more
common approach in addressing similar sized bands is
to eliminate questionable bands from the analysis in
addition to those confirmed as zooxanthellae in origin
(e.g. Goulet and Coffroth 1997, 2003a, b; Coffroth and
Lasker 1998).
Algal cultures provide the best source of clean algal
DNA with which to test putative cnidarian primers.
Culturing methods, however, tend to represent only a
Table 1 Cnidarian species used
in these studies
portion of the total genotypic variation attributed to
zooxanthellae that exist within a symbiotic cnidarian
(Goulet and Coffroth 1997; Rowan 1998; Santos et al.
2001; LaJeunesse 2002). Thus all possible zooxanthella
bands may not be represented in zooxanthella controls
derived from cultures and care should be taken to verify
that the algal culture that is used is representative of the
intact symbiosis. Because zooxanthella cultures isolated
from the cnidarian species of interest are often not
available, zooxanthellae from the host species can be
obtained via a tissue degradation technique (T. Wilcox,
personal communication; see below) in which the zooxanthellae remain viable while the host tissue and DNA
degrade over time. The remaining intact zooxanthellae
are representative of the in hospite zooxanthella population. Finally, zooxanthellae freshly isolated from the
host tissue, subjected to multiple washes and run
through a Percoll gradient have also been used as a
source of in hospite zooxanthella DNA (Tytler and
Davies 1983; Goulet and Coffroth 1997, 2003a, b;
Stochaj and Grossman 1997). However, as in the case of
the degradation protocol, this technique enriches for
zooxanthella DNA, but does not eliminate host DNA.
This technique, as well as the degradation protocol can
generate zooxanthella-biased DNA that can be used to
screen for specificity of cnidarian markers.
Materials and methods
Generation of cnidarian-specific molecular markers
A series of molecular markers were developed to
examine the genetic structure of cnidarian populations
and to infer phylogenetic relationships among scleractinian species (Table 1). Cnidarian samples were collected from Panama, the Flower Garden Banks, the
Florida Keys, Bermuda, and the Bahamas. Tissue was
preserved in either 95% ethanol, or salt-saturated 20%
dimethyl sulfoxide solution (Seutin et al. 1991), or frozen
in liquid nitrogen. DNA extractions of cnidarian tissue
followed Coffroth et al. (1992) or modifications of the
Prep-A-Gene DNA Purification Kit (BioRad). After
macerating the coral tissue (2–5 mm scraping of scleractinian tissue, approximately 5 mm of gorgonian tissue) in 200 ll grinding buffer [0.1 M NaCl, 0.1 M
EDTA (ethylenediaminetetraacetic acid), 0.05 M
Species
Taxonomic group
Tissue type
Symbiotic/asymbiotic
Molecular marker
M. cavernosa
Scleractinian
Sperm
Asymbiotic
P. astreoides
Scleractinian
Adult
Symbiotic
P. elisabethae
Gorgonian
P. kuna
Gorgonian
Adult
Larvae
Adult
Larvae
Symbiotic
Asymbiotic
Symbiotic
Asymbiotic
Sequencing
Microsatellites
Sequencing
Microsatellites
ISSRs
Microsatellites
Microsatellites
ISSRs
ISSRs
60
Tris–HCl; pH 8.0], the samples were incubated with
750 ll Binding Buffer (BioRad) at 65C for 10 min.
DNA bound to 15 ll Matrix (BioRad) during incubation at room temperature for 10 min, intermittently
mixing by inversion. The DNA-bound Matrix was pelleted via centrifugation at 14,000 rpm for 1 min and the
supernatant was discarded. The pellet was resuspended
with 750 ll Binding Buffer, centrifuged, and the supernatant was discarded. This was repeated two additional
times with Wash Buffer (BioRad). The pellet was dried,
resuspended in 50 ll Elution Buffer (BioRad), and
incubated at 65C for 10 min to elute the DNA from the
Matrix. After centrifugation, the supernatant containing
the DNA was transferred into a new tube, and the
remaining pellet was resuspended in 30 ll Elution Buffer, centrifuged, and the supernatant was added to the
first supernatant. To remove any remaining Matrix, the
supernatant was centrifuged and transferred to a new
tube twice. During the extraction, care was taken to
avoid rupturing zooxanthella cells in an effort to enrich
the template with host DNA (i.e. detergent was not included in grinding buffer and the tissue was not excessively homogenized). A DNA purification step using
polyethylene glycol, although not essential, increased the
number of successful PCR amplifications and was subsequently performed on each of the extracted scleractinian coral samples.
Cytochrome c oxidase subunit I In a study to infer
evolutionary relationships among Caribbean scleractinian corals (Shearer and Coffroth, in preparation), universal cytochrome c oxidase subunit I (COI) primers and
PCR conditions of Folmer et al. (1994) were utilized to
amplify a region of the COI gene from adults of 32 coral
species and 11 zooxanthella cultures (representing zooxanthella clades inhabiting these species; Table 2). The
presumed coral COI gene was sequenced in both directions using 5¢-IRD700 and IRD800 M13 labeled primers
(LI-COR Biotechnology Division, Lincoln, NE, USA)
and the SequiTherm EXCEL II DNA Sequencing KitLC (Epicentre Technologies, Madison, WI, USA)
according to manufacturers’ recommendations. Sequences were visualized on a 5.5% acrylamide gel using
a LI-COR’s NEN Global IR2 DNA Sequencer System
and scored using E-Seq version 1.1 (LI-COR Biotechnology Division, Lincoln, NE, USA). Sequences were
deposited into GenBank under accession numbers
AY451340 and AY451387. Nucleotide sequences were
aligned by eye and similarity to asymbiotic cnidarian
COI sequences deposited in GenBank was determined
using BLAST (Altschul et al. 1997).
Microsatellites To delineate population structure and
gene flow in corals, host-specific polymorphic microsatellite markers (di-, tri-, or tetra-nucleotide tandem
repeats dispersed throughout the genome) were successfully generated from sperm (Montastraea cavernosa),
asymbiotic larvae (Pseudopterogorgia elisabethae), and
adult tissue (Porites astreoides; Table 1) following the
enrichment protocol of Ciofi and Bruford (1998;
Gutiérrez-Rodrı́guez and Lasker 2004b; Shearer and
Coffroth 2004). Microsatellite loci were initially visualized on a 2% agarose gel stained with ethidium bromide
to determine if the PCR was successful. For each
Table 2 Source of zooxanthellae used to test various molecular markers
Clade A cultures
COI sequencing
ISSRs
Culture ID
Host
Culture ID
Host
Culture ID
Aiptasia pallida
FLap4a
FLap4b
Cass EL1
CassKB8
CassMJ300
FLCass
Pk708a
Pp719a
FLap4
CassMJ300
FLCass
CassEL1
Pk708a
Pp719a
Zs
FLap4
Cass EL1
CassKB8
CassMJ300
Pk708a
Pp719a
Zs
Aiptasia pallida
P. elisabethae
FLap3
Pe
Mastigias sp.
Mp
Aiptasia pallida
P. elisabethae
FLap3
Pe
Mastigias sp.
Montipora verricosa
Sinularia sp.
Mp
Mv
Sin
P. flexuosa
PurPflex
Aiptasia pallida
P. elisabethae
FLap3
Pe
Mastigias sp.
Montipora verricosa
Sinularia sp.
Mp
Mv
Sin
P. kuna
Pseudoplexaura porosa
Aiptasia pallida
Cassiopeia sp.
P. kuna
Pseudoplexaura porosa
Zoanthus sociatus
Aiptasia pallida
Cassiopeia sp.
P. kuna
Pseudoplexaura porosa
Zoanthus sociatus
Culture identification corresponds to the cultures listed in Santos et al. 2001
Zooxanthellae isolated from newly settled polyps
a
Clade C/F cultures
Host
Cassiopeia sp.
Microsatellites
Clade B cultures
61
microsatellite locus, PCR conditions were optimized to
adjust band intensity and reduce stutter bands. Microsatellite alleles were visualized on a 6.5% acrylamide gel
using a LI-COR’s NEN Global IR2 DNA Sequencer
System (LI-COR Biotechnology Division, Lincoln, NE,
USA) and scored with Gene ImagIR v3.55 (Scanalytics
Inc.). All polymorphic primers were tested against zooxanthella DNA to confirm that the markers were specific
to the cnidarian host. Even though M. cavernosa microsatellites were generated from asymbiotic sperm,
microsatellite primers were screened against cultures of
zooxanthella clade C, the dominant algal clade of this
species (Baker and Rowan 1997; LaJeunesse 2002), as
well as clades A, B and F algae to ascertain specificity to
host DNA (Table 2).
Pseudopterogorgia elisabethae microsatellite primers
were originally developed from adult tissue, however
screening these primers against zooxanthellae from an
algal culture isolated from P. elisabethae and against
asymbiotic P. elisabethae larvae, indicated that a high
proportion of these loci were specific for the zooxanthella DNA (Table 3). Analyses of chloroplast 23s
rDNA and ITS sequences of this zooxanthella culture
indicate this algae is representative of the numerically
dominant in hospite populations of this host species
(Santos et al. 2001). Microsatellite primers for this species were re-developed from asymbiotic larvae and tested
against the zooxanthella culture isolated from this host
species.
For P. astreoides, only symbiotic adult tissue was
available for microsatellite development. Primers were
screened against DNA from cultures of clades A, B and
C (Table 2), the predominant clades in P. astreoides
(Baker and Rowan 1997; LaJeunesse 2002) to determine
if zooxanthella DNA could be amplified. In addition,
the P. astreoides primers were also tested with zooxanthella DNA isolated from this species via the tissue
degradation protocols described below.
Inter-simple sequence repeats Intersimple sequence repeat (size-variable nucleotide regions between microsatellites amplified by a single primer; Wolfe et al. 1998)
banding patterns were generated from adult P. astreoides tissue and cultured zooxanthella clades A, B, and C
(Table 2) and from larvae and adult Plexaura kuna tissue to examine gene flow and population structure in
Table 3 The origin of polymorphic microsatellite loci generated
from various cnidarian tissues
Source
No. of
No. of
Polymorphic loci
clones
primers
sequenced designed Total No. of No. of
coral zoox
M. cavernosa sperm
P. elisabethae adults
P. elisabethae larvae
P. astreoides adults
31
19
27
42
a
13
11
10
21
The origin of two loci was not determined
5
6
2
7
5
0–2a
2
3
0
4–6a
0
4
these species. Each PCR reaction included 10% 10X
PCR buffer, 2 lM each dNTP, 11.7 mM MgCl2,
1.25 lM ISSR primer, 2.5 U Taq polymerase and
10–30 ng DNA template. Thermal cycling conditions
followed Wolfe et al. (1998). PCR products (25 ll) from
five different ISSR primers (17898-(CA)6RY; 17899(CA)6RG; 17901-(GT)6YR; M1-CAA(GA)5; 814.1(CT)8TG; Wolfe et al. 1998) were visualized on 2%
agarose gels stained with ethidium bromide. For each
coral DNA template, two independent PCR reactions
were run adjacent to each other, and only those bands
that were produced in the replicate lanes were scored.
Bands were scored as present or absent with the Kodak
1D (version 3.5.4). As ISSR primers are arbitrary and
based on microsatellite repeats randomly distributed
throughout the genome, zooxanthella DNA is expected
to amplify. Banding patterns of zooxanthellae are likely
to be dissimilar to coral patterns as it is unlikely that the
bands represent homologous regions of the two taxonomic groups.
Size-variable nucleotide region bands generated from
cultured zooxanthellae (clades A, B, and C; Table 2) and
degradation zooxanthella controls (from P. astreoides)
were compared to the banding patterns of several P.
astreoides samples to determine which bands may be
attributed to zooxanthella DNA. A comparison of
banding patterns between asymbiotic P. kuna larvae and
the maternal adult was used to determine the prevalence
of zooxanthella bands in the ISSR profiles from symbiotic host tissue.
Isolation of zooxanthella ‘‘control’’ DNA
DNA from zooxanthella cultures (Santos et al. 2001),
representing various algal clades (Rowan and Powers
1991), 23S rDNA chloroplast genotypes (Santos et al.
2002; Santos et al. 2003a), host species, and geographic
locations, was used as one type of zooxanthella control
(referred to as ‘‘culture controls’’) to determine the
source of the molecular markers (e.g. host or zooxanthellae). Because not all host species of interest had
corresponding zooxanthella cultures, zooxanthellae were
also obtained via degradation of the adult P. astreoides
tissue. In this technique (T. Wilcox, personal communication), the cnidarian tissue was macerated using a
tissue homogenizer and passed through a series of nylon
meshes (125, 70 and 20 lm) to remove host tissue. The
suspension was centrifuged at 800 rpm for 5 min and
the supernatant, containing tissue debris, was removed.
The pellet was resuspended in 10–50 ml filtered seawater
(0.2 lm, FSW), centrifuged and the supernatant discarded. This step was repeated and the pellet was
resuspended in 10–50 ml FSW and incubated overnight
at room temperature allowing any remaining host tissue
to breakdown. The suspension was centrifuged at
800 rpm for 5 min, the supernatant removed and the
pellet resuspended in 10 ml FSW. This process was repeated for 3–5 days until there was little debris visible
62
when the pellet was examined microscopically. Zooxanthella DNA extractions followed Coffroth et al.
(1992). DNA from this technique (referred to as ‘‘degraded controls’’) are representative of the in hospite
symbiont populations.
In P. astreoides, zooxanthella microsatellite primers
were inadvertently generated with the adult tissue as
four out of seven polymorphic loci (Table 3) were
amplified from clades A, B, C, and F zooxanthella cul-
Results
Cytochrome c oxidase subunit I The universal COI
primers amplified an approximately 700 bp PCR product from both coral and zooxanthella DNA, with
additional bands often present in zooxanthella samples
(Fig. 1). Sequences from coral tissue exhibited a high
sequence similarity (79%) with the asymbiotic sea
anemone, M. senile according to a BLAST search,
indicating that the sequences generated from adult tissue
were of coral origin. Thus, despite the fact that universal
primers are able to amplify presumably homologous
regions from coral and zooxanthellae, cnidarian DNA
appears to be preferentially amplified when both genomes are present, perhaps due to attempts to enrich the
template with host DNA.
Microsatellites For each species, only a small number
of polymorphic loci were successfully determined to be
coral in origin relative to the number of clones sequenced and primer sets designed (Table 3). Microsatellite primers designed from M. cavernosa sperm were
specific to the host and did not amplify zooxanthella
DNA (Fig. 2 and Table 3).
Initial attempts at generating microsatellite primers
from symbiotic adult tissue of P. elisabethae resulted in
80–100% of the primers amplifying cultured zooxanthellae from that host species (Table 3), while asymbiotic P. elisabethae larvae did not amplify (Fig. 3a). This
indicated that zooxanthella microsatellites were inadvertently generated from this tissue. All polymorphic
microsatellite loci generated from P. elisabethae larvae
were coral in origin (Table 3) as indicated by the fact
that the P. elisabethae zooxanthella culture did not
amplify whereas amplification of the asymbiotic larvae
was successful (Fig. 3b).
Fig. 1 Amplification of coral tissue using universal COI primers
resulted in a single product approximately 700 bp in size. DNA
from zooxanthella cultures resulted in an amplification product the
same size as that in coral tissue, often in addition to other bands
(indicated by arrows). Lanes 1–11 represent zooxanthella cultures
from clades A, B, C and F. Lanes 12 and 13 are amplifications from
adult P. astreoides and M. cavernosa, respectively, followed by a
negative control (N)
Fig. 2 Polymorphic coral microsatellite locus generated from
M. cavernosa sperm. Cultured zooxanthellae, lanes 1–13, representing various cultures and genotypes of clades A, B, C and F did
not amplify, while adult coral tissue produced one or two
polymorphic alleles (lanes 14–20). PCR reactions of zooxanthella
and coral samples were conducted simultaneously and the results
were reproducible. The negative control (N) did not produce an
amplification product. L 147 bp and 172 bp indicate molecular size
standard
Fig. 3 Microsatellite loci generated from P. elisabethae (a) adult
tissue and (b) asymbiotic larvae. Note that the locus generated from
adult tissue (a) did not amplify DNA from asymbiotic larvae (lanes
4–6) while the locus was amplified from the symbiotic adult DNA
templates (lanes 1–3) indicating an algal origin. This was supported
by amplification of the zooxanthella control (cz), a culture isolated
from P. elisabethae. The locus generated from asymbiotic larvae (b)
also amplified P. elisabethae adults (lanes 1–2) as well as asymbiotic
larvae (lanes 5–7). However, this locus did not amplify the
zooxanthella culture from this species (cz), indicating the locus
was host-specific. The positive control (P) was amplification of
clone DNA. Negative controls (N) did not produce an amplification product. Size standard is 100 bp DNA ladder
63
tures (Fig. 4a). Two polymorphic loci, however, were
unambiguously determined to be specific to P. astreoides
as zooxanthella cultures did not amplify, nor did the P.
astreoides zooxanthella obtained from the degradation
method (Fig. 4b). Primers for one P. astreoides locus
amplified coral samples and produced multiple bands
from a single zooxanthella culture, while other zooxanthella controls were negative (data not shown). Bands of
three host samples and this zooxanthella culture were
cloned and sequenced to determine if the primers were
amplifying homologous regions. The clones from the P.
astreoides samples exhibited identical sequences in the
region flanking the microsatellite while the zooxanthella
clones were not homologous and did not include a microsatellite repeat (data not shown), indicating that the
primers amplified a coral microsatellite locus and nonspecific regions of one zooxanthella culture.
Fig. 4 (a) Zooxanthella-specific and (b) host-specific polymorphic
microsatellite loci generated from P. astreoides adult tissue.
Zooxanthella loci were identified by amplification of zooxanthella
controls (a) lanes 1–7) which included various cultures and
genotypes of clades A, B and C. Microsatellite loci were determined
to be host-specific (b) if zooxanthella cultures (cz clades A, B and C;
lanes 1–9), degraded zooxanthella from adult P. astreoides (dz lanes
15–17) and negative controls (N) did not produce amplification
products. The positive control (P) was amplification of clone DNA.
L 122 bp, 147 bp and 172 bp indicate molecular size standard
Inter-simple sequence repeats Banding patterns of cultured zooxanthella controls were very different from
those of the adult P. astreoides tissue with only one band
potentially corresponding in size (Fig. 5a), indicating
most bands were cnidarian in origin. To make certain
that zooxanthella markers were not included in any
analyses, only bands specific to the host were used in the
data analyses, similarly sized host and symbiont bands
were eliminated. The majority of ISSR bands (10 out of
10 and 11 out of 14 bands from ISSR primers 17898 and
814.1, respectively) produced from maternal adult P.
kuna tissue were present in the asymbiotic larvae, indicating most bands were cnidarian in origin. Bands not
shared with larvae may either be attributed to inheritance from the paternal adult or to zooxanthella
Fig. 5 (a) Inter-simple sequence repeats patterns of P. astreoides
from adult tissue and zooxanthellae controls, including various
cultures and genotypes of clades A, B, C and F zooxanthellae.
Arrows indicate potential or actual zooxanthella bands in the adult
tissue samples, noting that there was little similarity between
zooxanthella and coral patterns. (b) ISSR patterns of two P. kuna
asymbiotic larvae (L1 and L2) and the maternal adult (run twice,
A1 and A2). A majority of the bands were cnidarian in nature as
indicated by bands shared between the mother and offspring.
Bands observed in the adult but not shared with larvae can be
attributed to polymorphism of the loci due to inheritance from the
paternal adult or to zooxanthella contamination. Size standard is
100 bp DNA ladder
64
contamination. Although cultured zooxanthella DNA is
successfully amplified with ISSR primers (Fig. 5a), as
with the COI sequence data, cnidarian DNA appears to
be preferentially amplified when both genomes are
present (Fig. 5b), perhaps due to attempts to enrich the
template with host DNA.
Discussion
Molecular genetic markers have proven to be powerful
tools in addressing evolutionary and ecological questions concerning reef corals, and these techniques will
become increasingly important as they are used to assess
population structure and address questions concerning
dispersal and connectivity, genetic basis for disease- and
bleaching-resistance and genetic implications of conservation practices. However, markers for the ‘‘wrong’’
genome can be generated rather easily and frequently.
Without testing for specificity, one may not realize that
the locus or marker in question is not from the species
of interest. Mistakenly including zooxanthella markers
in analyses for cnidarians will confound interpretations
of the results and lead to inappropriate conclusions as
biogeographic and phylogenetic patterns of zooxanthella do not necessarily reflect that of the host. For
example, microsatellite data suggests that population
structure of the symbiont does not necessarily mirror
that of the host species (Gutiérrez-Rodrı́guez 2003;
Santos et al. 2003b). In addition, co-evolution within
the cnidarian/zooxanthellae symbiosis has not been
established, as there is not a strong correlation between
host and symbiont phylogeny (Rowan and Powers 1991;
McNally et al. 1994; Baker and Rowan 1997). Thus,
verification that the molecular markers are cnidarian
in nature is critical because the outcome of these studies
is being put to practical use, data are being used to
answer questions concerning disease-and bleachingresistant coral strains, sources of recruitment and reef
connectivity and marine resource protection and
management.
The cost of interpreting inaccurate data is high;
however, it is relatively easy to overcome this hazard and
generate informative markers. As discussed above,
asymbiotic host tissue provides the best source of DNA
to generate markers. However, if uncontaminated DNA
is not available or markers based on arbitrary primers
are sought, one must rigorously screen the primers to
verify cnidarian specificity. One cannot assume a technique that provided informative markers in higher
organisms or non-symbiotic systems will automatically
yield markers for the genome of interest.
Contamination is also an obstacle for those developing molecular markers specific to zooxanthellae. As
zooxanthella cells exhibit few distinguishing morphological characteristics, genetic markers are developed to
clarify the taxonomy of the symbionts as well as to assess levels of genotypic diversity within and among
hosts. Despite difficulties obtaining zooxanthellae from
single-cell isolate cultures (Santos et al. 2001) and the
fact that cultures represent only a subset of the genotypes within a host (Goulet and Coffroth 1997; Rowan
1998; Santos et al. 2001), cultured zooxanthellae provide
the best source of clean DNA from which to generate
zooxanthella markers. As discussed above, using methods to physically separate symbionts and host can still
yield contamination of the algal DNA template with
residual coral DNA. Thus care should be taken to
ensure that zooxanthella markers generated from
zooxanthellae directly isolated from host tissue are algal
in origin.
The intimate association of host and algal DNA leads
to difficulties in analyzing micro- and macroevolutionary processes of cnidarian species. This study clearly
demonstrates that molecular markers intended to
investigate cnidarian genetic structure and evolution
were easily, frequently, and inadvertently generated
from the contaminating DNA of symbiotic algae.
Hence, modifications in methodology and/or rigorous
testing are required before interpretation of such results.
Those developing genetic markers for cnidarians generally understand such precautions and care is taken to
address this source of error (see references above).
Occasionally, however, attempts to confirm the origin of
the markers are not addressed or are not robust. This
leaves one to question the actual origin of the genetic
marker and the resultant data that have been generated.
Acknowledgments The authors would like to thank the Florida
Keys National Marine Sanctuary, the Flower Garden Banks National Marine Sanctuary and the Bahamas Department of Fisheries
for permission to collect and export scleractinian and gorgonian
samples. Special thanks to C. McNutt, S. Santos, R. Smith, and P.
Vize for collection and laboratory assistance, T. Wilcox for sharing
the degradation technique, the staff and scientists of the Don Gerace Research Center, Keys Marine Lab, Smithsonian Tropical
Research Institute and the National Undersea Research Center for
logistical support as well as the crews of the M/V Spree, M/V Sea
Ray and R/V Big Moe. We thank Aquarium of Niagara (Niagara
Falls, NY, USA) for access to synthetic seawater for culturing
zooxanthellae and Sherwood SCUBA for donating SCUBA
equipment for field collections. Thanks to J. Stamos for assistance
with figures. This research was supported by National Oceanographic and Atmospheric Administration’s National Undersea
Research Program (M.A.C.; 2000-15 and M.A.C. and J. Ault;
2002-12), the National Science Foundation (M.A.C.; OCE-9530057 and OCE-99-07319), the New York State Sea Grant Program (M.A.C. and H. Lasker; #R/XG-9), the National Undersea
Research Center at the Caribbean Marine Research Center
(M.A.C. and H. Lasker;#CMRC-99-3301 and 99-NRHL-01-01C)
and IIE/Conacyt/Fulbright-Garcı́a Robles Fellowship (C.G.R.).
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