Detection of microalgal resting cysts in European coastal sediments using a PCR-based assay

ARTICLE IN PRESS Deep-Sea Research II 57 (2010) 288–300

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Deep-Sea Research II journal homepage: www.elsevier.com/locate/dsr2

Detection of microalgal resting cysts in European coastal sediments using a PCR-based assay Antonella Penna a,, Cecilia Battocchi a, Esther Garce´s b, Silvia Angle s b, Emellina Cucchiari c, Cecilia Totti c, Anke Kremp d, Cecilia Satta e, Maria Grazia Giacobbe f, Isabel Bravo g, Mauro Bastianini h a

Department of Biomolecular Sciences, University of Urbino, Viale Trieste 296, 61100 Pesaro, Italy Institut de Cie ncies del Mar, CSIC, Passeig Marı´tim de la Barceloneta 37-43, 08003 Barcelona, Spain c Dipartimento di Scienze del Mare, Universita Politecnica delle Marche, Via Brecce Bianche, 60131 Ancona, Italy d Tv¨ arminne Zoological Station, University of Helsinki, 10900 Hanko, Finland e Dip. Botanica ed Ecologia Vegetale, University of Sassari, 07100 Sassari, Italy f Istituto per l’Ambiente Marino Costiero, CNR, Spianata S. Raineri 86, 98122 Messina, Italy g ˜ol de Oceanografı´a, Apdo. 1552, 36200 Vigo, Spain Instituto Espan h CNR-ISMAR, Istituto di Scienze Marine, Castello 1364/A, 30122 Venezia, Italy b

a r t i c l e in f o

a b s t r a c t

Available online 23 September 2009

A PCR-based assay was developed and applied to sediment and sediment trap samples for the detection of different cysts belonging to dinoflagellates and raphidophytes in European coastal areas. Oligonucleotide primers were designed based on the ITS-5.8S and LSU ribosomal gene sequences. The specificity and sensitivity of the PCR assay were assessed using genomic DNA from clonal cultures, plasmid copy number of cloned target sequences, as well as from sediment samples. Qualitative PCR determinations of different cysts in sediment and sediment trap samples were compared to taxonomic examinations by light microscopy. This molecular methodology permitted a fast and specific detection of target cysts in sediment samples. We also detected dinoflagellate and raphidophyte cysts at concentrations not detectable by microscopic methods or that are difficult to identify. The results given by molecular and microscopic methods were comparable. However, higher values of positive detection for target cysts were obtained by PCR than with microscopy. Some taxa were detected in 100% of the samples using PCR, while others were only found in 10% of the samples. The data obtained in this study showed that the PCR-based method is a valid tool for cyst identification in marine sediments. & 2009 Elsevier Ltd. All rights reserved.

Keywords: HAB species Mediterranean Sea PCR Ribosomal genes Resting stages Sediments

1. Introduction Harmful Algal Blooms (HABs) are recurring events in European coastal waters (Giacobbe et al., 2007; Smayda, 2007). Many HAB species are responsible for these events and the majority are dinoflagellates (IOC Taxonomic reference list, http://www.bi.ku. dk/ioc/). Most HAB dinoflagellates display heteromorphic life cycles, including motile planktonic stages, as well as immotile benthic resting cysts. All stages of the life cycle, and especially the dormant cysts, have a large impact on bloom dynamics. The bloom development of cyst-forming HAB species may also be dependent on the presence of cyst beds seeding local blooms (Garce´s et al., 2002; Steidinger and Garce´s, 2006). While great effort has been dedicated to the study of the planktonic stages of these microorganisms, we still have limited knowledge of their other life stages, especially benthic stages. In some dinoflagellates

 Corresponding author.

E-mail address: [email protected] (A. Penna). 0967-0645/$ - see front matter & 2009 Elsevier Ltd. All rights reserved. doi:10.1016/j.dsr2.2009.09.010

and raphidophytes, the resting stages with their resistant walls permits the species to tolerate environmental conditions and therefore to expand their geographical distribution (Amorim et al., 2001; McGillicuddy et al., 2003; De Boer et al., 2004; Edvardsen and Imai, 2006). Knowledge of the cyst bed composition and location in bottom sediments can provide information on the long-term presence of a species in an area, as well as enabling prediction of subsequent blooms. Mapping the cyst distribution of HAB species also provides information on cyst transport to new areas, depending on oceanic currents. Moreover, cyst distribution reflects the sedimentary dynamics and the location of blooms in the overlying surface waters (Angle s et al., 2010). Further, cyst assemblage data provide an indication of the potential plankton diversity reservoir in a locality. All this information is necessary to understand and predict potential HAB development and expansion. Therefore, monitoring the distribution of cyst densities in coastal areas prior to an outbreak is important for localizing hot spots for blooms and to minimize the damage caused by harmful blooms (Bravo et al., 2006).

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Studies on resting cyst diversity and distribution have been hampered by difficulties in the reliable identification of speciesspecific resting stages, since dinoflagellate and raphidophyte cysts do not always have species-specific morphological features. Traditional microscopic methods, which are commonly employed for monitoring of HAB species, often do not allow identification, unless the corresponding vegetative form can be provided through in vitro germination of the respective cysts. Molecular techniques and in particular, PCR techniques based on the amplification of targeted ribosomal genes, have been developed for the rapid and accurate detection and quantification of vegetative cells of HAB species both in culture and field samples (Galluzzi et al., 2004; Coyne et al., 2005; Godhe et al., 2007; Penna et al., 2007). The PCR detection technique can be applied to samples from sediment cores and sediment traps containing a variety of cysts. Such molecular-based assays can accurately and rapidly identify a variety of specific taxa in the sediments, overcoming the problem of taxonomic identification by microscopy (Bolch, 2001; Bowers et al., 2006). However, to date, only a few dinoflagellates have been identified in sediment samples using the PCR approach (Godhe et al., 2002; Kamikawa et al., 2007). The success of the PCR-based approach in sediments is strictly dependent on the inhibitory substances contained in the sediment material (Saito et al., 2002). Different strategies are applied to overcome the problem of PCR inhibition caused by contaminating substances: the use of commercial nucleic acid extraction kits that remove inhibitors and facilitate purification; the use of thermostable DNA polymerase; the dilution of template DNA prior to PCR assay and the use of BSA (bovine serum albumin) in the PCR assay. In this study, a PCR-based assay was developed and applied to sediment and sediment trap samples for the detection of resting cysts from several taxa. The specificity and sensitivity of the PCR assay were assessed both in clonal microalgal culture and field sediment samples that were collected from several European coastal areas. Oligonucleotide primers specific to several HAB dinoflagellate species, namely Alexandrium spp., Lingulodinium polyedrum, Protoceratium reticulatum, Gymnodinium catenatum, G. nolleri, and the raphidophyte Fibrocapsa japonica were applied in the PCR-based assay to identify the corresponding cyst morphotypes in sediment and sediment trap samples from different areas of the Mediterranean, Baltic Sea, and the East Atlantic coast. The assay was based on the LSU, 5.8S rRNA genes and ITS regions as target regions for the taxa specific primers and was validated on the genomic DNA of the clonal cultures and preserved cyst samples. A protocol for species-specific PCR detection of cysts was developed based on the use of a nucleic acid purification kit. The specificity and sensitivity of each assay was determined and comparisons of the qualitative determinations of the PCR analysis and optical microscopy were made. Data on the detection of the resting cyst stage of the different HA species are shown.

2. Methods 2.1. Study sites Surface sediment samples were collected during the years 2006 and 2007 from 38 sampling stations located at 11 different sites (Table 1) in the Mediterranean and Baltic Sea and the East Atlantic (Fig. 1). In the Mediterranean Sea, the sampling sites were distributed in four regional Seas: the North West Adriatic, Ionian, Tyrrhenian, and Catalan Sea. The sampling sites in the Baltic Sea and East Atlantic Ocean were located in the coastal Northern Baltic Sea and the western coast of Spain, respectively. Stations were located in areas where dinoflagellate or raphidophyte

289

blooms occur, and in sediment accumulation areas, as indicated by deposition maps, e.g. for the Arenys harbor (Garce´s et al., 2004; Angle s et al., 2010) and Ria de Vigo (Bravo and Anderson, 1994).

2.2. Cyst sample collection, purification, and microscopic morphotype identification Vertical sediment samples up to 8–10 cm in depth were collected using a gravity corer (7.2 cm internal diameter). Three replicate samples were taken per station. Sediment samples of Arenys de Mar (Catalan Sea) and Olbia (Tyrrhenian Sea) harbors were collected and processed as indicated in Satta et al. (2010). Sediment samples (7.5 ml) were extracted from the undisturbed 3-cm surface layer in the sample core container using a syringe and were transferred into a 50-ml conical tube. In addition, sediment samples of 10 ml were collected from the bottom 3 cm of the sediment surface layer by a SCUBA diver using a plastic syringe. Samples were preserved by adding Lugol’s solution (Andersen and Throndsen, 2003) in filtered seawater and storing them in the dark at 4 1C until analysis. Three samples were taken from sediment traps, collected at the Arenys harbor (Catalan Sea) and Baiona site (East Atlantic coast); the sampling of sediment traps was carried out following the method of Garce´s et al. (2004). Settled material was collected from the traps every 4 days and trap subsamples of 48 ml were fixed with Lugol’s solution and kept in the dark at 4 1C until analysis. Subsamples of 2.5 ml of suspended sediments were sonicated (Bandelin, Germany) to disaggregate cysts from sediment particles, sieved on steel membrane (Endecotts, UK) using size fractionation of 100- and 10-mm membranes and transferred into filtered seawater. The density gradient method was applied to sediment subsamples to separate cysts from detrital material (Amorim et al., 2001; Bravo et al., 2006). Samples were then used both for microscopic examination (2 ml subsamples) and molecular analysis (5-ml subsamples). The resting cysts were identified and counted under an inverted microscope at 200  and 400  ¨ magnification by scanning the entire Utermohl sedimentation chamber. Different morphotypes of dinoflagellate and raphidophyte cysts were classified according to shape, color, wall thickness, and size. Cyst abundance was expressed as the cyst number per volume of wet sediment sample and total cyst number processed for molecular analysis.

2.3. Genomic DNA extraction of sediment and trap samples Sediment samples (5 ml), obtained as above, were centrifuged at 4000 rpm for 10 min at room temperature; the supernatant was gently discarded and 20 ml of filtered seawater were added; samples were again centrifuged at 4000 rpm for 10 min at room temperature. This washing step was repeated twice; then, the cyst pellet was resuspended in 1 ml of sterile MilliQ water and centrifuged at 10,000 rpm for 5 min at room temperature; the supernatant was discarded and the pellet was frozen at 80 1C until nucleic acid extraction. The pellet was thawed at + 65 1C for 15 min, and re-frozen at  80 1C for 15 min; this freeze–thaw step was repeated twice. After the last thawing, the pellet was sonicated for 20 min in an ultrasonic bath and then added to a tube containing beads and lysis buffer of the UltraClean Soil DNa Kit (MoBio Lab In., Solana Beach, CA). DNA extraction and purification were carried out according to the manufacturer’s instructions. Purified genomic DNA was quantified on an agarose gel using serially diluted Lambda DNA Marker (MBI Fermentas, Germany) and a gel-documentation apparatus (Bio-Rad, Hercules, CA, USA).

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Table 1 Sediment samples collected along several coastal areas in the Mediterranean, Baltic Sea and Eastern Atlantic, cyst abundance per sample and processed cyst abundance in the molecular based assay. Total processed cyst abundance

Station coordinates

Sampling period

Station number

Sampling type

Cyst abundance (ml  1  wet sediment)

NW Adriatic Sea, Italy NW Adriatic Sea, Italy NW Adriatic Sea, Italy NW Adriatic Sea, Italy NW Adriatic Sea, Italy NW Adriatic Sea, Italy NW Adriatic Sea, Italy NW Adriatic Sea, Italy NW Adriatic Sea, Italy NW Adriatic Sea, Italy NW Adriatic Sea, Italy NW Adriatic Sea, Italy NW Adriatic Sea, Italy N Tyrrhenian Sea, Italy N Tyrrhenian Sea, Italy N Tyrrhenian Sea, Italy S Tyrrhenian Sea, Italy Ionian Sea, Italy Ionian Sea, Italy Ionian Sea, Italy Ionian Sea, Italy N Catalan Sea, Spain N Catalan Sea, Spain N Catalan Sea, Spain N Catalan Sea, Spain

Venezia, C10 Ancona, South A Ancona, South C Ancona, South B Ancona, South-out CD Ancona, North-out AB Ancona, South-in AB Ancona, North C Ancona, North-in ABL Ancona, North-out ABL Ancona, North B Ancona, North A Ancona, North-out DER Olbia A Olbia B Olbia G Vulcano St.1C, Siracusa St. 2C, Siracusa St. E, Siracusa Aquacult. B, Siracusa Arenys, St. 15 Arenys, St. 11 Arenys, St. 23-1 Arenys, St.12 + St. 17

January 2006 July 2006 July 2006 July 2006 December 2006 December 2006 December 2006 December 2006 December 2006 December 2006 July 2006 July, 2006 July 2006 May 2007 May 2007 May 2007 May 2006 March 2007 March 2007 March 2007 March 2007 April 2006 April 2006 June 2006 June 2006

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25

Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Sediment core Trap sample Trap sample Sediment core Sediment core

428 126 94 77 120 59 78 306 170 247 512 470 386 121 59 97 7 159 134 1 56 36 38 1857 2656

4286 1890 1415 1105 1805 890 1175 4590 2555 3700 7680 7050 5790 375 177 291 100 365 281 3 175 1807 1895 3714 10640

N Catalan Sea, Spain

Arenys, St. 8 + St. 13

June 2006

26

Sediment core

2591

10364

N Catalan Sea, Spain N Catalan Sea, Spain N Catalan Sea, Spain N Catalan Sea, Spain East Atlantic, Spain East Atlantic, Spain East Atlantic, Spain East Atlantic, Spain East Atlantic, Spain Baltic Sea, Sweden Baltic Sea, Finland Baltic Sea, Finland

Arenys, 2 Arenys, 1/3 Arenys, 2 - 4 Arenys, 1/5/6 Baiona 2 Ria de Vigo a Ria de Vigo b Baiona 28 Baiona 6 ¨ Himmersfjarden Storfjarden ˚ ¨ Langsk ar

451 150 431 300 431 300 431 300 431 300 431 300 431 300 431 410 431 420 431 430 431 430 431 420 431 410 401 550 401 550 401 550 381 250 371 030 371 030 371 030 371 020 411 340 411 340 411 340 411 340 340 6000 411 340 340 6600 411 340 411 340 411 340 411 340 421 070 421 130 421 130 421 070 421 070 581 590 591 510 591 470

October 2006 October 2006 December 2006 December 2006 August 2006 March 2006 March 2006 July 2006 December 2006 March 2007 May, 2007 May 2007

27 28 29 30 31 32 33 34 35 36 37 38

Sediment core Sediment core Sediment core Sediment core Trap sample Sediment core Sediment core Trap sample Sediment core Sediment core Sediment core Sediment core

2415 2345 2837 3132 1140 15 114 655 280 434 8801 6342

4830 9380 11348 18792 5700 180 1368 3600 1400 39060 792135 285390

0000 N, 121 450 6000 E 0900 N, 131 410 0000 E 0000 N, 131 410 0700 E 0500 N, 131 410 0300 E 0000 N, 131 410 0700 E 0900 N, 131 410 0000 E 0500 N, 131 410 0300 E 0400 N, 131 230 0000 E 0100 N, 131 220 0000 E 0400 N, 131 190 0000 E 0400 N, 131 190 0000 E 0100 N, 131 220 0000 E 0400 N, 131 230 0000 E 4500 N, 91 300 2500 E 4500 N, 91 300 4500 E 1500 N, 91 300 1400 E 0700 N, 141 570 1100 E 5100 N, 151170 0100 E 2100 N, 151170 4000 E 5100 N, 151 170 01 E 0900 N, 151 170 20 E 6900 N, 21 330 6800 E 6900 N, 21 330 6800 E 7300 N, 21 330 3400 E 7200 N, 21 330 7000 E 411 N, 21 330 4400 E 6900 N, 21 330 4200 E 411 N, 21 330 7000 E 7300 N, 21 330 3400 E 7300 N, 21 330 3400 E 7300 N, 21 330 3400 E 7300 N, 21 330 3400 E 1200 N, 81 500 1500 E 0800 N, 81 510 1400 E 0800 N, 81 510 1400 E 1200 N, 81 500 1500 E 1200 N, 81 500 1500 E 0700 N, 171 430 6000 E 31400 N, 231130 00100 E 33400 N, 231; 200 28400 E

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Localities and sampling stations

A. Penna et al. / Deep-Sea Research II 57 (2010) 288–300

Geographical area

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Fig. 1. Locations of the sampling stations in the Mediterranean Sea, East Atlantic and Baltic Sea (see Table 1 for locality names).

Table 2 Clonal cultures used together to test the specificity of genus and species specific PCR identification assay. Species

Strain

Collection site

Source

Alexandrium minutum Chaetoceros socialis Fibrocapsa japonica Gymnodinium catenatum Gymnodinium nolleri Scrippsiella hangoei Scrippsiella rotunda Scrippsiella trochoidea Prymnesium parvum Skeletonema marinoi Ostreopsis ovata Pseudo-nitzschia spp.

VGO663 CCMP 204 CBA-1 GC12V 922I SHTV-1 CBA-4 VGOS3V CCMP708 CBA-2 CNR-D1 CBA-4

Tyrrhenian Sea, Olbia, Italy Catalan Sea, Marseille, France Adriatic Sea, Ancona, Italy East Atlantic, Vigo, Spain Kattegat, Denmark ¨ Baltic Sea, Tvarmine, Finland Adriatic Sea, Pesaro, Italy East Atlantic, Vigo, Spain North Atlantic, Scotland, UK Adriatic Sea, Pesaro, Italy Ligurian Sea, Genova, Italy Adriatic Sea, Pesaro, Italy

Fraga S. Berland B. Totti C. Figueroa R.I. Ellegard M. Kremp, A. Ingarao C. Bravo I. Droop M. Penna A. Giacobbe M.G. Capellacci S.

2.4. Microalgal cultures and genomic DNA extraction Clonal cultures were used for taxon-specific PCR-based assays (Table 2). Cultures were kept in F/2 or L1 media (https://ccmp. bigelow.org/) at 20 71 1C in a 12:12 h (light:dark) photoperiod. Illumination was provided by photon irradiance of 100 mmol m–2 s–1. Culture subsamples containing 100,000 cells of each target species were used for the genomic DNA extraction using a DNeasy Plant Kit (Qiagen, Valencia, CA, USA) according to Penna et al. (2005).

2.5. PCR amplification detection assay Genus (Alexandrium) and species-specific (Alexandrium minutum, A. taylori, A. tamarense, A. catenella, L. polyedrum, P. reticulatum, F. japonica) primers were designed in the 5.8S rDNA-ITS regions and/or were derived from Penna et al. (2007). The primers for the genus Scrippsiella and species G. catenatum, G. nolleri and Scrippsiella hangoei were designed in the LSU and 5.8S rDNA-ITS regions by using OLIGO software ver. 6.65 and BioEdit software ver. 7.0.5 (Table 3). The BLAST (Basic Local Alignment Search Tool) analysis of alignment nucleotide sequences (Altschul et al., 1990) were as follows: primer G. catenatum: F0 ATTGCAGAATTCCGTGAA, total alignment length: 18 nt, overall identity: 100% (18/18 nt). Primer G. nolleri F0 TTTCAGCGATGGATGTCT, total alignment length: 18 nt, overall identity: 100% (18/18 nt). Primer Scrippsiella sp. F0 CTGAAA

GGAAAGCGAATGGAG, total alignment length: 22 nt, overall identity: 63.64% (14/22 nt). Primer S. hangoei F0 TTCGT TCGGAAGTGGTTT, total alignment length: 18 nt, overall identity: 100% (18/18 nt). The LSU and 5.8S rDNA-ITS sequences for G. catenatum VGO-744 (AM998536), G. nolleri VGO-663 (AM998534), G. nolleri 922I (AM998535), Scrippsiella rotunda CBA-4 (AM998538), and S. trochoidea VGO-S3V (AM998537) were obtained in this study; the 5.8S rDNA-ITS sequences for S. hangoei were obtained from GenBank. Amplification from sediment and sediment trap samples for the detection of different cyst taxa was performed in an Applied Biosystems DNA Thermo Cycler 2720 (Foster City, CA, USA). PCR amplifications were carried out directly using genus- or species-specific primers or by following two steps as follows: an initial PCR using eukaryotic primers targeting the ITS-5.8S or LSU ribosomal genes; then, a second (i.e. nested) PCR reaction, using genusand species-specific primers on amplified products of ribosomal genes (Fig. 2).

2.6. Direct PCR-based assay Direct PCR using genus- or species-specific primers was as follows: reaction tubes contained a 50 mL mixture of 200–400 mM of each dNTP; 0.2–0.4 mM of each primer; 3.0–6.0 mM MgCl2; 1  Reaction Buffer (Eppendorf, Germany); 0.5–1  Enhancer Buffer (Eppendorf, Germany); 0.2–1.5 mg/ul BSA (Table 4); 0.5 U of Taq Polymerase (Eppendorf, Germany) and 0.5–1.0 ng of sediment

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Table 3 Oligonucleotide primers targeting the ITS-5.8S rDNA and LSU regions of different phytoplankton genera and species used in this study. Target taxa

Forward Primer

Reverse primer

Forward primer sequence (50 - 30 ) Reverse primer sequence (50 - 30 ) ’

Amplification size (bp)

G + C%

Primer positions

Alexandrium spp.a

5.8S-50

5.8S-30

135

A. catenellaa

ITS1c

5.8S-30

A. minutum

ITS1m

5.8S-30

A. tamarensea

5.8S-50

ITS2t

A. tayloria

ITS1t

5.8S-30

Fibrocapsa F

Fibrocapsa R

G. catenatumb

Gymno-cat F

Gymno-cat R

G. nollerib

Gymno-nolleri F

Gymno-nolleri R

L. Poly GF

L. Poly GR

P. reticulatum

P. ret F

P. ret. R

Scrippsiella sp.b

Scripp F

Scripp R

S. hangoeib

Scripp-hangoei F

Scripp-hangoei F

F0 -GCAADGAATGTCTTAGCTCAA R0 -GCAMACCTTCAAGMATATCCC F0 -AGCATGATTTGTTTTTCAAGC R0 -GCAMACCTTCAAGMATATCCC F0 -CATGCTGCTGTGTTGATGACC R0 -GCAMACCTTCAAGMATATCCC F0 - TGTTACTTGTACCTTTGGGA R0 - ACAACACCCAGGTTCAAT F0 -TGGTGTTTGAATGCGGTTGT R0 -GCAMACCTTCAAGMATATCCC F0 -GCAGAGTCCAGCGAGTCATCA R0 -TAATATCCCAGACCACGCCAGA F0 -ATTGCAGAATTCCGTGAA R0 -GATCGATGCGAATGAAAC F0 -TTTCAGCGATGGATGTCT R0 -TGAAGGCACGATTGACAC F0 -ATGTGTTCTCATCGGATGTTG R0 -CACAGTACCGCTGCCACTTAAA F0 -TGCTGATTGCCATCTATCTT R0 -CAGAAGCGCGTTAAACAG F0 -CTGAAAGGAAAGCGAATGGAG R0 -CCGCAAATGAGTTCCAACAAG F0 -TTCGTTCGGAAGTGGTTT R0 -CCCGAGAGCACCTTAACA

38.0 42.8 33.3 42.8 52.3 42.8 40.0 44.4 45.0 42.8 57.1 50.0 38.9 44.4 44.4 50.0 45.5 50.0 40.0 50.0 47.6 47.6 44.4 55.6

5.8S (50 -30 ) 5.8S (30 ’50 ) ITS1 (50 -30 ) 5.8S (30 ’50 ) ITS1 (50 -30 ) 5.8S (30 ’50 ) 5.8S (50 -30 ) ITS2 (30 ’50 ) ITS1 (50 -30 ) 5.8S (30 ’50 ) 5.8S (50 -30 ) ITS2 (30 ’50 ) 5.8S (50 -30 ) ITS2 (30 ’50 ) 5.8S (50 -30 ) ITS2 (30 ’50 ) ITS1 (50 -30 ) ITS2 (30 ’50 ) ITS1 (50 -30 ) ITS2 (30 ’50 ) LSU (D1-D2) LSU (D1-D2) ITS1 (50 -30 ) ITS2 (30 ’50 )

a

F. japonica

a

L. polyedrum

a

a

Degenerate Degenerate Degenerate Degenerate a b

code code code code

226 212 134 297 180 290 350 383 382 221 274

D = A/G/T. M = A/C. Y = C/T. R = A/G.

From Penna et al. (2007). designed in this study.

Fig. 2. Schematic representation of the PCR assays based on the use of genus and species-specific or eukaryotic primers. Direct or nested PCR assays were used for the PCR amplification of target taxa cyst in sediment and trap samples.

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Table 4 Different concentrations of dNTPs, primers, MgCl2, Enhancer and BSA specific for each microalgal taxon used in the PCR based assay. Taxa

dNTPs (mM)

Primers (mM)

MgCl2 (mM)

Enhancera

BSA (mg/ml)

Alexandrium spp. Alexandrium catenella Alexandrium minutum Alexandrium tamarense Alexandrium taylori Gymnodinium catenatum Gymnodinium nolleri Fibrocapsa japonica Lingulodinium polyedrum Protoceratium reticulatum Scrippsiella spp. Scrippsiella hangoei

0.2 0.2 0.2 0.4 0.4 0.4 0.4 0.4 0.4 0.4 0.1 0.4

0.4 0.4 0.4 0.4 0.4 0.4 0.4 0.2 0.2 0.4 0.4 0.4

6.0 6.0 6.0 6.0 6.0 6.0 6.0 3.0 3.0 3.0 6.0 3.0

No added 0.5 1.0  No added 0.5 1x No added No added 0.5 No added 0.5 No added

0.2 0.2 1.0 1.0 0.4 1.5 0.4 0.2 No added 0.2 0.2 0.4

The Taq Master PCR enhancer is a buffer additive that improves thermostability (enzyme half-life) and processing of Taq DNA polymerase by stabilizing the enzyme during PCR. a

The TaqMaster PCR enhancer is a buffer furnished with the Taq Polymerase (Eppendorf, Germany).

template DNA. PCR conditions were as follows: an initial denaturation step of 10 min at 95 1C, 40 cycles of 30 s at 95 1C, 30 s at 58 1C, and 30 s at 72 1C, and a final extension step of 7 min at 72 1C. 2.7. Nested PCR-based assay For the nested PCR eukaryotic-specific primers targeting the ITS-5.8S rDNA (Adachi et al., 1994) and LSU rDNA (Scholin et al., 1994) primers were used in the first PCR. Reaction tubes contained a 25 mL mixture of: 200 mM of each dNTP; 2 pmol of each primer; 4 mM MgCl2; 1  Reaction Buffer (Eppendorf, Germany); 0.5 U of Taq Polymerase (Eppendorf, Germany) and 0.5–1.0 ng of sediment template DNA. PCR conditions were as follows: an initial denaturation step of 10 min at 95 1C, 40 cycles of 30 s at 95 1C, 30 s at 55 1C and 30 s at 72 1C, and a final extension step of 7 min at 72 1C. The PCR-amplified products, which were derived from the PCR assay using universal eukaryotic primers and were not directly visualized on agarose gel, were used as template in nested PCR amplification with genus- or species-specific primers using 1 mL of the first PCR-amplified product; the PCR conditions were as described above for the direct PCR amplification with the exception of 35 cycles rather than 40. 2.8. Sequencing The 5.8S gene and ITS regions of genomic DNA from microalgal species were amplified using ITSA/ITSB primers (Adachi et al., 1994). The PCR-amplified products were purified using the Qiagen Purification Gel Extraction Kit (Qiagen, CA, California) according to the manufacturer’s instructions and were then directly sequenced. PCR-amplified products were sequenced on an ABI PRISM 310 Genetic Analyser (Applied Biosystems, USA) using the dye terminator method according to the manufacturer’s instructions (ABI PRISM Big Dye Terminator Cycle Sequencing Ready Reaction Kit, Applied Biosystems). Genus- and species-specific PCR-amplified products obtained from sediment samples, such as those from Station 23 (Arenys) for Alexandrium, Station 11 (Ancona, North B) for A. minutum, Station 8 (Ancona, North C) for A. tamarense, Station 27 (Arenys) for Scrippsiella, Station 18 (Siracusa) for L. polyedrum, Station 12 (Ancona, North A) for P. reticulatum, Station 9 (Ancona, North-in ABL) for F. japonica, Station 33 (Ria de Vigo) for G. catenatum and Station 31 (Baiona) for G. nolleri, were excised from the gel, purified using QIAquick Gel extraction Kit (Qiagen) and directly sequenced.

2.9. Specificity and sensitivity of the PCR assay The specificity and sensitivity of the PCR reactions had already been assessed for Alexandrium, A. minutum, A. tamarense, L. polyedrum, P. reticulatum and F. japonica in Penna et al. (2007). The specificity of the Scrippsiella spp., S. hangoei, G. catenatum and G. nolleri oligonucleotide primers were tested by amplifying target DNA of the various microalgal genera and species from both clonal cultures of target species and sediment samples with designed species-specific primers. The Scrippsiella spp., S. hangoei, G. catenatum and G. nolleri genomic DNA was amplified with specific primers in the presence of non-target mixed genomic DNA obtained from different clonal strains; 1 ng of genomic DNA was used for each background species. The 5.8S rDNA-ITS regions of Scrippsiella spp., S. hangoei, G. catenatum and G. nolleri were amplified and cloned into the Pcr 2.1 vector following the manufacturer’s instructions (Invitrogen, Carlsbad, CA, USA). Plasmids containing the target ITS5.8S and LSU rDNA sequences and non-target mixed genomic DNA were used as a positive and negative control, respectively. To assess the sensitivity of the PCR assay, 100, 10 and 1 pg of each genomic DNA of Scrippsiella spp., S. hangoei, G. catenatum and G. nolleri were spiked into 10, 5 and 1 ng of the mixed cyst population DNA, providing background sediment DNA in the mixture, which was then subjected to the PCR assays. The sensitivity assays were done in triplicate for each amount of genomic DNA and taxon examined. The background sediment sample was checked for the absence of the target taxa (Scrippsiella spp., S. hangoei, G. catenatum and G. nolleri) by microscopy. Assessment of sensitivity was also performed on a retroviral cloned sequence in an external (non-microalgal) plasmid DNA (Casabianca et al., 1998); this non-microalgal plasmid DNA was spiked as 104, 103, 102 and 101 copies into 10, 5 and 1 ng of mixed cyst population DNA. The PCR products were resolved on a 1.8% (w/v) agarose, 0.5X TBE buffer gel and were visualized by standard ethidium bromide staining under UV light.

3. Results 3.1. Sample collection and cyst content identification by microscopy A total of 38 sediment and sediment trap samples were collected during the study in several coastal localities of the Mediterranean and Baltic Sea and the north-western coast of Spain. A relatively high diversity of cyst taxa was found, with a total of 32 cyst morphotypes identified by microscopy (Table 5). Dinoflagellate

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Table 5 List of cyst morphotypes from sediment and sediment trap samples of the Mediterranean and Baltic Sea and East Atlantic coast sampling stations identified by microscopy. Taxa

Dinophyceae Gymnodiniales Polykrikos spp. Polykrikos kofoidii Polykrikos schwartzii Gymnodinium spp. G. catenatum G. nolleri G. impudicum Gonyaulacales Alexandrium spp. Alexandrium minutum Alexandrium pseudogoniaulax Alexandrium ostenfeldii Gonyaulax sp. G. spinifera Woloszynskia halophila Lingulodinium polyedrum Protoceratium reticulatum Peridiniella catenata Peridiniales Pentapharsodinium dalei Pentapharsodinium tyrrhenicum Protoperidinium spp. Protoperidinium cfr. claudicans Protoperidinium conicum Protoperidinium compressum Protoperidinium cfr. divaricatum Protoperidinium granii Protoperidinium cfr. oblongum Scrippsiella spp. Scrippsiella hangoei Scrippsiella lachrymosa Scrippsiella trochoidea Undetermined dinoflagellates Raphidophyceae Fibrocapsa japonica Total cyst type

Adriatic Sea

Ionian Sea

Tyrrhenian Sea

Catalan Sea

+

+

Baltic Sea

+ + +

+ + +

+ +

+

+ +

+

+ +

+ +

+ +

+

+ +

+ +

+

+ +

+ + +

+

+

+

+ +

+ + + +

+

+

+

+ + +

ribosomal genes of one dinoflagellate genus and three species gave amplified fragments of different base pair lengths. PCRamplified products for the genus Scrippsiella were 221 bp long and for the three species, S. hangoei, G. catenatum and G. nolleri, ranged from 274 to 350 bp in length. The specificity of the genus- and species-specific primers was assessed both by multiple alignment on the silico platform BLAST and by PCR amplification of the genomic DNA from each genus and species in the presence of mixed non-target taxa clonal strain DNA together with target species clonal strains. The target genus- and species-specific primers showed high specificity in all selected PCR-based assays and no other detectable bands were observed; non-target genomic DNA used as background DNA templates was not amplified. The specificity of the PCR-based assay for the other dinoflagellate genera and species, and the raphidophyte F. japonica, was assessed in an earlier study by Penna et al. (2007). The sensitivity and the absence of inhibitors of the PCR-based assay using specific primers were assessed on the plasmid target cloned sequence of a murine retroviral complex. The PCR assay on plasmid DNA was sensitive enough to detect ten copies of the cloned sequence in the presence of 1 ng of sediment DNA as background. The sensitivity of the PCR assay was also assessed using genomic DNA as a template. The sensitivity of the PCR-based assay carried out on genomic DNA corresponded to the specific PCR amplification of 1 pg. The presence of 1 ng of background genomic DNA from the sediment sample, containing mixed dinoflagellate resting cysts, did not have an effect on the sensitivity of any of the species-specific PCR-based assays. Assays using genomic DNA were inhibited by Z5 ng of the background DNA from the sediment sample (data not shown).

+

3.3. Detection of dinoflagellate and raphidophyte resting cysts in coastal sediments

+ +

+

+

+ + +

+

+

+

+

+

+

+ + +

+

+

+ + +

+ 18

15

12

14

12

+ +

species belonging to Peridiniales and Gonyaulacales dominated the assemblages at all stations. In particular, Protoperidinium and Scrippsiella species within the Peridiniales were dominant; but cysts of potentially toxic dinoflagellate species, such as Alexandrium spp., A. minutum, and Gymnodinium spinifera, were also recorded. Gymnodinium cysts were also found, but were not identified at the species level with the exception of G. catenatum, G. nolleri and Gymnodinium impudicum. Further, F. japonica cysts were identified in sediment samples from the northern Adriatic Sea. 3.2. Specificity and sensitivity of the PCR-based assay PCR amplifications using primers designed in the highly variable ITS regions and conserved 5.8S and LSU (D1/D2)

PCR amplification was performed on several Lugol-fixed sediment and sediment trap samples collected during the years 2006 and 2007 along the coastal areas of the Mediterranean Sea, north-western coast of Spain and Baltic Sea to detect the presence of either dinoflagellate resting cysts belonging to the genera Alexandrium and Scrippsiella and species A. minutum, A. tamarense, A. catenella, A. taylori, G. catenatum, G. nolleri, L. polyedrum, P. reticulatum, and S. hangoei or raphidophyte resting cysts belonging to the species F. japonica (Table 6). These samples contained mixed cyst communities including the target morphotypes, with total cyst abundances ranging from 3.0  101 to 7.9  105. The amount of target taxa in the analyzed samples ranged from 1.0  101 to 2.3  105 of cysts. Several direct PCR amplifications, using genus- and species-specific primers on sediment samples, which contained low numbers of target cysts, resulted in faint bands or the absence of bands on agarose gel. Thus, nested PCR amplification reactions were performed on sediment and trap samples using genus- and species-specific primers. Nested PCR amplification reactions with eukaryoticspecific primers targeting the 5.8S-ITS and LSU rDNA regions first, and then amplifying the genus- or species-specific regions, always gave positive PCR products from the genomic DNA in sediment samples. PCR products of the expected size for each genus and species were detectable in the samples containing the target cyst morphotype, thus confirming the ability to amplify the target genomic DNA by direct or nested PCR-based assay. The PCR assay detected the presence of the resting stages belonging to different dinoflagellate taxa even if target cysts were not observed in the sediment and sediment trap samples by microscopy examinations. The PCR assays were positive for the presence of cysts of the genera Alexandrium and Scrippsiella and the species

Table 6. PCR based assay on the ITS-5.8S and LSU rDNA and microscopy analysis of various phytoplankton genera and species processed cyst number from sediment and trap samples collected during the period 2006-2007 along several coastal areas of the Mediterranean, Baltic Sea and East Atlantic. Station number andregional Sea

A. tamarense

Scrippsiella spp.

Processed cyst number

Species specific PCR assay

Processed cyst number

Species specific PCR assay

Processed cyst number

Genus specific PCR assay

Processed cyst number

+ +a + +a + +a +a + +a + + + + a a + + +a +a a a + + + + + + + +a + + + + + +a + + +

393 225 350 265 270 65 140 715 445 612 1010 890 825 3 n.i. n.i. n.i. 23 98 1 6 23,684 20,455 825 859 967 744 494 326 437 5700 n.i. n.i. 3600 n.i. 1300 650 1750

+ +a + +a + + a +a + + a  + +  a a +  +a +a a a + + + + + + + +a + + +a +a + +a +  

393 n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. 3 n.i. n.i. n.i. 23 98 n.i. n.i. 23,684 20,455 825 859 967 744 494 326 437 5700 n.i. n.i. 3600 n.i. 300 n.i. n.i.

a a a a a a a +  + a   +  a  a  + a a a a a a a a a a a a a a a a a a a a a

n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i.

+ + + + + + + + + + + + + + a + a + a n.a. +a +a a + + + a + + + +a a + +a +a +a + + + + +

74 180 260 210 300 230 180 965 580 917 1882 1515 805 141 87 59 n.i. 45 58 n.i. 77 14,211 15,682 712 1203 1118 1286 1229 1251 1429 n.i. n.i. n.i. n.i. n.i. 7830 2970 5820

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Genus specific PCR assay

295

1, NW Adriatic 2, NW Adriatic 3, NW Adriatic 4, NW Adriatic 5, NW Adriatic 6, NW Adriatic 7, NW Adriatic 8, NW Adriatic 9, NW Adriatic 10, NW Adriatic 11, NW Adriatic 12, NW Adriatic 13, NW Adriatic 14, N Tyrrhenian 15, N Tyrrhenian 16, N Tyrrhenian 17, S Tyrrhenian 18, Ionian 19, Ionian 20, Ionian 21, Ionian 22, Catalan 23, Catalan 24, Catalan 25, Catalan 26, Catalan 27, Catalan 28, Catalan 29, Catalan 30, Catalan 31, East Atlantic 32, East Atlantic 33, East Atlantic 34, East Atlantic 35, East Atlantic 36, Baltic Sea 37, Baltic Sea 38, Baltic Sea

A. minutum

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St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St.

Alexandrium spp.

P. reticulatum

F. japonica

G. catenatum

G. nolleri

Species specific PCR Processed cyst assay number

Species specific PCR Processed cyst assay number

Species specific PCR Processed cyst assay number

Species specific PCR Processed cyst assay number

Species specific PCR Processed cyst assay number

+ +a +a +a +a +a +a + + + +a + +    n.a. + +a +a +a   a +a a a a a +a a a a a +a a a a

+ a a a a a +a +a +a +a +a +a +a a a a n.a. a a a a + a a a a a a a a a +a +a a a a a a

a a a a a a a +a +a +a + + +a a a a n.a. a a a a   a a a a a a a a a a a a a a a

a a a a a a a a a a a a a a +a a n.a. a a a a a a a a a a a a a +a + + a + a a a

a a a a a a a a a a a a a a a a n.a. a a a a a a a a a a a a a +a a a   a a a

607 100 80 15 70 50 75 415 220 395 175 480 200 n.i. n.i. n.i. n.i 216 58 n.i. 58 n.i. n.i. n.i. 10 n.i. 7 n.i. n.i. 11 n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i.

n.i. n.i. n.i. n.i. n.i. 5.0 10 60 40 52 37 30 50 n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i.

n.i. n.i. n.i. n.i. n.i. n.i. n.i. 85 80 165 305 255 120 n.i. n.i. n.i. n.i n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i.

n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. 180 1368 n.i. 1400 n.i. n.i. n.i.

n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i. n.i.

The PCR detections of the species A. catenella, A. taylori and S. hangoei in the sediments were not shown. A. catenella and A. taylori were not present in the sediment samples. S. hangoei was present only in the Baltic Sea sediment samples. + = positive amplification;  = negative amplification. n.i.= not identified; n.a. = not available. a

Nested on the ITS-5.8S, LSU rDNA.

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1, NW Adriatic 2, NW Adriatic 3, NW Adriatic 4, NW Adriatic 5, NW Adriatic 6, NW Adriatic 7, NW Adriatic 8, NW Adriatic 9, NW Adriatic 10, NW Adriatic 11, NW Adriatic 12, NW Adriatic 13, NW Adriatic 14, N Tyrrhenian 15, N Tyrrhenian 16, N Tyrrhenian 17, S Tyrrhenian 18, Ionian 19, Ionian 20, Ionian 21, Ionian 22, Catalan 23, Catalan 24, Catalan 25, Catalan 26, Catalan 27, Catalan 28, Catalan 29, Catalan 30, Catalan 31, East Atlantic 32, East Atlantic 33, East Atlantic 34, East Atlantic 35, East Atlantic 36, Baltic Sea 37, Baltic Sea 38, Baltic Sea

L. polyedrum

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St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St. St.

296

Station number andregional Sea

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297

Fig. 3. Comparison of the total sample number processed both by PCR and microscopy methods for the detection of different taxa in sediment and trap samples collected at different regional Seas of the Mediterranean and Baltic Sea and the East Atlantic coast during the years 2006 and 2007. Analyses of positive and negative PCR amplifications compared with the corresponding positive and negative microscopic examinations of the sediment samples are shown.

A. minutum, A. tamarense, G. catenatum, G. nolleri, L. polyedrum, P. reticulatum, and S. hangoei (data not shown). The resting cysts of species A. catenella and A. taylori were never detected in the sediment samples examined by PCR assay and microscopy (data not shown). The positive detection of the two genera and eight species obtained by PCR assay and microscopy were compared (Fig. 3). For F. japonica, 12/12 samples were positive for both methods; in contrast, the species-specific identification of A. tamarense, G. catenatum and G. nolleri was confirmed by the molecular method only. The number of positive detections obtained by PCR assay was higher than microscopy determinations in 14 samples for A. minutum, 5 samples for Alexandrium spp. and Scrippsiella spp., 4 samples for P. reticulatum, and 2 samples for L. polyedrum. Furthermore, both methods revealed negative detections of target taxa cysts in the field samples, when the total number of processed samples was considered (Fig. 3). These values were higher for Gymnodinium species and F. japonica, while lower values of negative detections were found for the Gonyaulacales and Peridiniales groups. False negatives were also obtained by the PCR assay in some cases, in which cysts were positively identified by microscopy.

4. Discussion 4.1. Molecular techniques Molecular methodologies are highly specific, sensitive, and rapid techniques for the diagnostic identification of microbial eukaryotes in aquatic environments (Galluzzi et al., 2005, 2008; Gescher et al., 2008). Moreover, gene amplification techniques (PCR methods) with taxon-specific oligonucleotide primers and probes have been extensively developed and have shown great potential with regard to the identification and enumeration of many harmful dinoflagellate species (Dyhrman et al., 2006; Handy et al., 2006). The PCR technique has mostly been utilized for the identification of vegetative cells in the water column rather than other life cycle stages, such as the resting stages in sediments, with the exception of Godhe et al (2002) and Erdner et al. (2010).

The goal of this study was to evaluate the utility of the PCRbased technique for the identification of dinoflagellate and raphidophyte cysts in coastal sediment samples compared to microscopy determinations. Furthermore, this study proposes providing an assessment of the PCR method used, since the application of the PCR method to sediments is problematic due to inhibitory contaminants and the low concentrations of target cysts. Based on the results obtained in this study, the technique produced higher detection efficiency than the microscopic method, as shown by the higher positive percentage identifications of the target harmful cysts in sediments and sediment traps. 4.2. PCR-based analysis The PCR-based assay using new and tested oligonucleotide primers was developed to identify several dinoflagellate taxa and the raphidophyte F. japonica in sediments and sediment traps. The PCR assay for genus- and species-specific cyst detection was performed in mixed culture samples and then validated in cyst sediment samples using different concentrations of PCR components for each assessed target genus- or species-specific primers. It was found that BSA can co-precipitate the inhibitor compounds and that the TaqMaster Enhancer buffer makes the Taq DNA Polymerase less sensitive to exogenous PCR-inhibiting contaminants. The ribosomal genes were useful in designing genusspecific Scrippsiella and species-specific G. catenatum, G. nolleri, and S. hangoei primers; the highly variable ITS regions with more conserved rDNA genes permits discrimination at the inter-species level in sediment samples containing various cyst populations, as demonstrated for other genera and species in seawater samples in our previous study (Penna et al., 2007). 4.3. Sensitivity of the PCR assay The sensitivity of the PCR assay was assessed by spiking plasmid DNA with a known amount of sediment genomic DNA as background. The sensitivity of the PCR for amplifying the lowest amount of target DNA was determined by the amplification of ten copies of plasmid DNA/ng background sediment DNA. Furthermore, the PCR assay amplified 1 pg of genomic DNA from cultured Scrippsiella and Gymnodinium species in the presence of 1 ng of mixed cyst population DNA as background,

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while the taxon-specific PCR-based assay was inhibited in the presence of 10 and 5 ng of sediment genomic DNA as background. It is likely that the presence of some inhibitory substances, which are not completely eliminated from sediments during extraction and purification using the commercial kit, can negatively affect the PCR reaction. Inhibitor substances, such as humic acids, polyphenols, polysaccharides and metals, and nuclease activity, are the major concern when extracting genomic DNA from marine sediments (Stults et al., 2001). In fact, the coprecipitation of compounds that inhibit PCR confuses the molecular analyses of field samples by producing false negative results (Tebbe and Vahjen, 1993). The most common strategy used to overcome PCR inhibition by contaminants is dilution to a lower concentration of the template DNA. In this study, we applied a total DNA extraction and purifying procedures using a commercial kit to eliminate the potential inhibitors of the PCR reaction.

4.4. Specificity of PCR assay The PCR-based assay was effective for the qualitative detection of the target cysts in sediment and sediment trap samples. Total DNA extraction was accomplished with a commercial kit that was effective in obtaining the total quantity of DNA visible on agarose gel, which was then quantifiable for subsequent PCR reactions. The commercial kit was also successful in removing major inhibitors, since the direct PCR amplification using either taxonspecific or eukaryotic-specific primers targeting the LSU or 5.8S rDNA and ITS was positive. Nevertheless, absence of PCR amplifications were also obtained as direct PCR using genusand species-specific primers or eukaryotic primers often resulted in very faint bands or no observed bands at all, thus yielding apparent false negatives. A nested taxon-specific PCR amplification of the first amplified fragment using eukaryotic-specific primers, always gave a positive PCR reaction. The lack of PCR-amplified fragments was observed in those sediment samples containing a low fraction of target taxa cysts (n = 7) (Fig. 3). Direct and nested PCR-based amplifications with isolated single cysts have been achieved by minimizing cyst DNA loss by the Bolch (2001) and Ki and Han (2007) methods. However, a suitable method for the rapid and specific detection of dinoflagellate cysts in numerous sediment samples during monitoring activity is required. The single cyst PCR technique requires the manipulation and isolation of the single cysts from the sediment samples under a microscope, and a single isolated cyst may be sufficient to carry out only a few PCR amplification reactions.

4.5. Molecular PCR and microscopic determinations of sediment samples PCR detection of the target species was compared with the microscopic analyses of the same sediment and sediment trap samples. The molecular technique provided a higher positive detection rate of target cysts than microscopy. Target cysts were detected by PCR in sediment samples presenting mixed resting stages at low concentrations or concentrations below the detection limit of the microscopy method, and when it was not possible to recognize the target cyst taxonomically. Based on the PCR-based assay higher rates of positive detection were observed for the genera Alexandrium and Scrippsiella, and species A. minutum, A. tamarense, G. catenatum, G. nolleri, L. polyedrum and P. reticulatum due to a widespread distribution. The raphidophyte F. japonica cysts were identified equally in sediment samples by the two methods. PCR analysis detected 100% of A. tamarense, G. nolleri and G. catenatum cysts in sediment samples. This is a relevant finding as the different morphotype cysts of Gymnodinium spp. and Alexandrium spp. are quite difficult to recognize and distinguish (Bolch et al., 1999; Bravo et al., 2006). Furthermore, when PCR and microscopy methods were applied to sediments and seawater for the detection of target taxa (resting or vegetative stages), the PCR method produced a higher number of positive detections compared to the positive microscopy determinations both in sediment and seawater substrates (Fig. 4). The PCR-based assay detected the presence of A. minutum and A. tamarense in the Mediterranean Sea, where the two species were retrieved from both sediments and seawaters. The PCR assay also detected cysts of Alexandrium spp. in all three stations sampled in the Baltic Sea. The PCR assay also confirmed the presence of A. minutum cysts, which were identified by microscopy in Himmerfj¨arden sediments from the Swedish eastern coast. This is the first record of this species in the lower saline waters in the northern areas of the Baltic Sea, implying that A. minutum might be expanding and potentially cause harmful algal blooms in this area. Further, in the present study, S. hangoei and Gymnodinium sp. cysts were detected in the Baltic sediment samples by the PCR detection assay, confirming the tentative identification of the S. hangoei cyst morphotype in the sediment samples. While Woloszinskia halophila and Gymnodinium sp. form characteristic cysts, which can be recognized by microscopy (Kremp et al., 2005), S. hangoei cysts lack distinctive features and the morphotype might easily be confused with other smooth, transparent and colorless cyst types. In the sediment samples of the Baltic Sea, overall cyst numbers were approximately ten times higher than in the sediment samples from the East Atlantic and

140 120 100

y = 1.270x + 4.111

80

R2 = 0.947

60 40 y = 0.925x + 3.282 R2 = 0.945

20 0

0

20

40

60 80 100 Samples positives by microscopy

120

140

Fig. 4. Number of samples that showed positive detection with both methods (PCR-based assay and microscopy) for different dinoflagellate and raphidophyte cysts (’) and vegetative cells (E), in sediments and seawater samples, respectively. The diagonal line (1:1) indicates equal detection of both methods in sediments and seawater samples. Sediment samples were collected in European coastal waters during 2006 and 2007. Positive PCR and microscopy determinations on seawater samples were derived from Penna et al. (2007).

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the Mediterranean Sea. Such high total cyst concentrations are due to the dominance of W. halophila, which encysts in large numbers after the spring bloom, resulting in exceptionally high cyst fluxes in the sediments (Kremp and Heiskanen, 1999). The genus Scrippsiella was also highly abundant among the cystforming dinoflagellates in the examined areas; Scrippsiella cells are commonly found in coastal waters of these regions (Montresor et al., 1994). The life strategy of this genus includes a short dormancy period with rapid turnover rates in cyst/vegetative cells, which may explain the formation of the bloom and its abundance along with its long-term presence in a widespread area. The identification of Gymnodinium cysts was quite difficult due to the many almost indistinguishable morphotypes, some of which correspond to the same species (Matsuoka and Fukuyo, 2003). Within the genus Gymnodinium, G. catenatum is a toxic species producing a reticulated cyst, which makes it easier to distinguish from other cysts, but it is difficult to differentiate from G. nolleri cysts (Bolch et al., 1999). This species is reported to be recurrent on the western Atlantic coast of Spain (Bravo and Anderson, 1994) and in the Alboran Sea in the Mediterranean Sea (Bravo et al., 1990). Moreover, it has appeared sporadically in some regions of the western Mediterranean Sea and the cysts were detected during occasional surveys in southern western Mediterranean sediments (Calbet et al., 2002). In this study, G. catenatum cysts were found in the Spanish Atlantic sediments by microscopy and PCR assay, and also by the PCR method in one locality of the Tyrrhenian Sea where the cysts had never been detected by microscopy. With regards to the raphidophyte species, reports on F. japonica cysts are found in the in vitro study of De Boer et al. (2004), and the identification of the cyst morphotype is limited to marine sediments (Yoshimatsu, 1987). In the Mediterranean Sea, F. japonica blooms have only been registered in the Tyrrhenian and north-western Adriatic Seas (Cucchiari et al., 2008). In this latter coastal area, F. japonica produces abundant blooms during the summer period (Totti C., pers. comm.). In the present study, cysts of F. japonica were detected in the north-western Adriatic sediment samples by both methods, proving for the first time the presence of F. japonica resting stages in the area where this species causes blooms. Moreover, the taxonomic identification of cyst morphotype by microscopy was confirmed by molecular assay. The cysts of the toxic species L. polyedrum and P. reticulatum were found at almost all sampling stations, since they are widespread in European coastal seawaters.

5. Conclusions In this study the specificity and sensitivity of the PCR-based technique for the detection of target cysts in marine sediment and sediment trap samples was demonstrated. The PCR method permitted higher detection efficiency than the microscopic method, illustrated by the higher positive percentage identification of the harmful target cysts in sediments. Knowledge of species composition is important to understand bloom events in the coastal areas; it is also crucial to have information on the presence of novel and potentially introduced taxa and to confirm the recurrent events of a species. In the future, the PCR method could be used for mapping the distribution of target species cysts in coastal sediments, particularly given its high specificity and sensitivity.

Acknowledgments We thank S. Casabianca and A. Casabianca for molecular analysis assistance and suggestions; S. Fraga for culture strains;

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S. Capellacci for technical assistance. Thanks to the two anonymous reviewers who made an effort in improving the paper. This study was financed by the EU funded Research Project SEED (GOCE-CT-2005-003875). E. Garce´s was sustained by a Ramon y Cajal contract from the Spanish Ministry of Science and Education.

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