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Please cite this article as: Rundberget, T., Gustad, E., Samdal, I.A., Sandvik, M., Miles, C.O. AConvenient and Cost-Effective Method for Monitoring Marine Algal Toxins with PassiveSamplers, Toxicon (2009), doi: 10.1016/j.toxicon.2009.01.010

This is a PDF file of an unedited manuscript that has been accepted for publication. As aservice to our customers we are providing this early version of the manuscript. Themanuscript will undergo copyediting, typesetting, and review of the resulting proof before it ispublished in its final form. Please note that during the production process errors may bediscovered which could affect the content, and all legal disclaimers that apply to the journalpertain.

Accepted Manuscript

Title: A Convenient and Cost-Effective Method for Monitoring MarineAlgal Toxins with Passive Samplers

Authors: Thomas Rundberget, Eli Gustad, Ingunn A. Samdal, Morten

Sandvik, Christopher O. Miles

PII: S0041-0101(09)00046-4

DOI: 10.1016/j.toxicon.2009.01.010

Reference: TOXCON 3404

To appear in: Toxicon

Received Date: 3 July 2008

Revised Date: 6 January 2009Accepted Date: 16 January 2009

http://dx.doi.org/10.1016/j.toxicon.2009.01.010

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A Convenient and Cost-Effective Method for Monitoring Marine Algal

Toxins with Passive Samplers

Thomas Rundberget1, Eli Gustad

2, Ingunn A. Samdal

1, Morten Sandvik

1,

Christopher O. Miles1,3

1National Veterinary Institute, PB 8156 Dep., NO-0033 Oslo, Norway

2

Institute of Marine Research, Fldevigen Research Station, Fldevigen, N-4817 His, Norway

3AgResearch Ltd., Ruakura Research Centre, Private Bag 3123, Hamilton, New Zealand

*Corresponding author: National Veterinary Institute

Tel: +47 2321-6231; Fax: +47 2321-6201

E-mail address: thomas.rundberget@vetinst.no

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Abstract

Passive sampling disks were developed based on the method of MacKenzie et al. (2004) and

protocols were formulated for recovering toxins from the adsorbent resin via elution from

small columns. The disks were used in field studies to monitor in situ toxin dynamics during

mixed algal blooms at Fldevigen in Norway. Examples are given from time-integrated

sampling using the disks followed by extraction and high performance liquid

chromatography-mass spectrometry (HPLC-MS) analysis for azaspiracids, okadaic acid

analogues, pectenotoxins, yessotoxins and spirolides. Profiles of accumulated toxins in the

disks and toxin profiles in blue mussels ( Mytilus edulis) were compared with the relative

abundance of toxin-producing algal species. Results obtained showed that passive sampling

disks correlate with the toxin profiles in shellfish. The passive sampling disks were cheap to

produce and convenient to use and, when combined with HPLC-MS or enzyme-linked

immunosorbent assay (ELISA) analysis, provides detailed time-averaged information on the

profile of lipophilic toxin analogues in the water. Passive sampling is therefore a useful tool

for monitoring the exposure of shellfish to the toxigenic algae of concern in northern Europe.

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Keywords:Dinophysis, okadaic acid, dinophysistoxin, azaspiracid, passive sampling,

shellfish toxin, algal toxin

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1. Introduction

Over the last decade there has been an increase in the commercial cultivation and exploitation

of shellfish along the Norwegian coast. Contamination of shellfish with biotoxins from micro-

algae can be a problem for public health not only in Norway, but world wide (Hallegraeff,

1993; Toyofuku, 2006; Camacho et al., 2007), and many countries regulate the biotoxins in

shellfish (FAO/WHO/IOC, 2005). The Norwegian marine biotoxin monitoring programme

involves phytoplankton identification and enumeration, together with analysis of shellfish

flesh. The Norwegian Food Safety Authorities have a public surveillance program for algal

toxins in mussels. During the 2007/2008 season the algal monitoring was performed weekly

while chemical analysis of shellfish was performed monthly, from February to December and

only at selected places (3540 locations), and the programme is not able to cover all of the

vast Norwegian coastline.

Analysis of biotoxins in the shellfish flesh is required to determine the safety of the product

for consumption. However, analysis of shellfish is time consuming, technically demanding

and expensive, so it is not ideal as a tool for monitoring the progress of toxigenic blooms. In

addition, many of the toxins are metabolised in shellfish during digestion and assimilation,

and the increased variety and complexity of the metabolite profile makes toxin quantification

even more challenging. Phytoplankton monitoring involves collecting a concentrated sample

of the algae, shipping the sample to a suitable laboratory, and then enumerating the

identifiable toxigenic species (Lund et al., 1958). Phytoplankton monitoring has the ability to

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monitoring and shellfish analysis has historically provided a reasonable degree of protection

to shellfish consumers in Norway and elsewhere (Hallegraeff, 1993; van Egmond et al., 1993;

Batoreu et al., 2005).

Recently, alternatives have been sought to improve marine biotoxin monitoring. Of these,

passive sampling methods have shown much promise as tools for measuring aqueous

concentrations of a wide range of priority pollutants. The first passive sampling methods were

aimed at monitoring the concentrations of dissolved inorganic compounds in surface water

(Benes and Steinnes, 1974). Since then, there has been a rapid development in the use of

passive sampling devices (Huckins et al., 1990; Sodergren, 1990; Alvarez et al., 2004). Some

of the general features of different passive sampling devices have previously been reviewed

(Vrana et al., 2005; Stuer-Lauridsen, 2005). In comparison to traditional water sampling,

passive samplers offer the ability to integratively sample a range of environmental

contaminants over an exposure period, mimicking biological uptake while potentially

avoiding the heterogeneity and clean-up problems implicit with biological matrices (Verhaar

et al., 1995; Kot-Wasik et al., 2007). Recently, MacKenzie et al. (2004) introduced the idea of

monitoring algal toxins by passively adsorbing them directly from seawater using solid-phase

adsorbents. These so-called solid-phase adsorption toxin tracking (SPATT) devices,

consisting of bags sewn from polyester mesh containing activated polystyrydivinylbenzene

resin, adsorb lipophilic algal toxins dissolved in seawater. The SPATT bags provide a more

convenient means to perform time-averaged sampling prior to, or during, algal blooms than

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present in the sample extracts. Also, since the devices adsorb toxins released directly from the

algae into the water, the toxin profile is much simpler than the metabolite profile usually

found in shellfish. This results in easier assays, fewer toxins to quantify, and lower detection

limits for the targeted toxins. The resin used in the SPATT bags was tested and validated by

MacKenzie et al. (2004) for a range of algal toxins found in New Zealand (pectenotoxin-2

(PTX-2), PTX-2 seco acid (PTX-2 SA), yessotoxin (YTX), ocadaic acid (OA) and

dinophysistoxin-1 (DTX-1)).

The suitability of this approach for monitoring algal toxins in Norwegian waters was

investigated. As part of the study, the practicality of the device was improved by introducing a

frame in which the HP-20 resin is restrained to form a passive sampling disk. This design is

simple, cheap, more easily assembled and disassembled than the sewn SPATT bags, and is

well suited to high throughput processing of samples. In the trials, results obtained from

analysis of the passive sampling disks were compared to those from shellfish analyses and

phytoplankton monitoring at Fldevigen in Norway. Because passive sampling devices

containing the HP-20 resin have been validated for analysis of PTXs, YTX, OA/DTXs and

azaspiracids (AZAs) (MacKenzie et al., 2004; Fux et al., 2008), no attempt was made to

perform validation in the present investigation. Parts of the work have been reported in a

preliminary form in an earlier communication (Rundberget et al., 2006).

2. Materials and Methods

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2004a), PTX-2 SA (Miles et al., 2004b), PTX-2 (Miles et al., 2004c), OA, DTX-1 and DTX-2

(Larsen et al., 2007) and a semi purified mixture of AZA-1, -2 and -3 (unpublished) were

available in our laboratory from previous work.

2.2 Passive sampling disks.

Passive samplers were constructed from 100-m nylon mesh (Sefar AG, Heiden, Switzerland)

folded in half, a 75 mm diameter plastic embroidery frame (Permin, Copenhagen, Denmark)

and HP-20 resin (DIAION HP-20, Mitsubishi Chemical Corporation, Tokyo, Japan). The

resin (3.0 g) was placed between the two layers of nylon mash, and clamped tightly in the

embroidery frame so as to form a thin layer of resin between the layers of mesh. A No. 2

fishing swivel (Mustad, Gjvik, Norway) was attached to the outer ring of the embroidery

frame to provide a point of attachment during deployment (Figure 1). The resin was activated

by soaking the packed disk in methanol for 15 min and washing in deionised water, as

described in the resin-manufacturers instructions. The activated passive sampling disks were

placed in an air-tight plastic bags and stored cold (but not below 0 C) prior to and after

deployment in the sea.

2.3 Extraction of toxins from disks.

The embroidery ring was opened, and the used resin was quantitatively transferred to a 25 mL

Varian Bond-elute reservoir fitted with a 20 m nylon frit (Varian, Palo Alto, CA) and

washed free of salts with 3050 mL deionized water. Excess water was drawn from the

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dryness in vacuo. The residue was dissolved in 1.0 mL 80% MeOH, centrifuged, and the

supernatant analyzed by HPLC-MS. Alkaline hydrolysis was performed by mixing 200 L of

5 M NaOH with 0.8 mL methanolic HP-20 extract. The mixture was left to react at 37 C for

45 min, followed by addition of 210 L of 5 M HCl. Samples were filtered through 0.2 m

Spin-X filters prior to chromatographic analysis.

2.4 Field trials.

Trials were performed at the Marine Research Institute, Fldevigen, on the south-west coast

of Norway. Passive sampling disks were taken from their packaging and deployed by

attaching them to a fixed point at 1 m depth, and leaving them for the required time. The disks

were then rinsed briefly with fresh tap water, sealed in an air-tight plastic bag, and shipped to

the laboratory for analysis. Simultaneously, shellfish were harvested weekly and kept at 20

C prior to analysis, while algal cell counting was performed 3 times weekly.

2.5 Shellfish samples.

Frozen samples of blue mussels (Mytilus edulis) were thawed, and the flesh was removed

from the shells and homogenized using an Ultra Turrax

homogenizer (IKA

, Werke GmbH

& Co. KG, Staufen, Germany). The homogenates were stored at 20 C until extracted.

Homogenized shellfish (2 g) was extracted three times with 6 mL methanol by vortex mixing

for 2 min, and centrifuged at 2500 g for 5 min between extractions. The three extracts were

combined in a 20 mL volumetric flask, and the volume was adjusted to 20 mL with methanol.

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2.6 Algal cell counts.

Phytoplankton is routinely monitored in Fldevigen Bay three times per week. Every

Monday, Wednesday, and Friday, samples for enumeration and identification of

phytoplankton were taken as an integrated sample using a flexible hose, from 03 m depth,

from the same location as mussels and passive samplers. The water sample was preserved

using neutral Lugols solution. Smaller flagellates and algae in high concentration were

counted under the light microscope using a PalmerMaloney chamber (200 magnification),

with a detection limit of 104

cells/L (Palmer and Maloney, 1954). Larger dinoflagellates were

counted on semitransparent filters according to the description of Fournier (1978).

Examination in light microscope (100 magnification) was performed on 50 mL of the

sample that was gently filtered onto the filter for cell-counting, giving a detection limit of 20

cells/L.

2.7 HPLC-MS analysis.

Liquid chromatography was performed on a Symmetry C18 column (3 m, 50 2.1 mm)

(Waters, Milford, MA) using a Waters 2670 HPLC module. Separation was achieved by

linear gradient elution, starting from acetonitrilewater (35:65 v/v, both containing 5 mM

ammonium formate and 0.01% formic acid) rising to 100% acetonitrile over 10 min, held for

5 min, then switched back to the start-eluent. The HPLC system was coupled to a Quattro

Ultima Pt triple quadrupole mass spectrometer operating with an electrospray ionization (ESI)

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collision energy settings were optimized while continuously infusing (syringe pump) 20

ng/mL of the toxin standards at 3 L/min. Detection of the analytes was performed by

multiple reaction monitoring (MRM) in either positive (AZA-1 842.5>672.5, AZA-2

856.5>672.5, AZA-3 828.5>658.5, PTX-2 876.5>823.5, PTX-2 SA 894.5>823.5, PTX-12

874.5>821.5, 20-methylSPX-G/SPX-C 706.5>164.2) or negative (OA/DTX-2 803.5>255.1,

DTX-1 817.5>255.1, YTX 1141.5>1061.5) ionization mode. Except for PTX-12 and 20-

methylSPX-G/SPX-C, and DTX-2, which were quantified from calibration curves of PTX-2

and OA, respectively, all toxins were quantified using external calibration curves of standard

specimens dissolved in 80% MeOH.

3. Results and discussion

3.1 Practical aspects of the improved disks

The HP-20 resin used in the disks has been tested and validated for a range of lipophilic

biotoxins by others (MacKenzie et al., 2004; Fux et al., 2008), and no attempts were made to

perform validation in this study. The main improvement over the SPATT bags of MacKenzie

et al. (2004) lies in the design of the frame in which the HP-20 resin is retained. This design

simplifies the preparation of the activated disks, their deployments and the subsequent toxin

extraction compared to the sewn SPATT bags. The new design was quick and easy to use, the

frames and algal mesh could be washed and reused, and the frames hold the resin in a

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water in much the same way as mussels might. However, the disks have the advantage that

there is no toxin metabolism, they are more easily stored and cheaper to transport, and provide

a much cheaper and cleaner extract for the analytical laboratory.

The adsorption rate of lipophilic toxins from sea water by HP-20 resin is fast. MacKenzie et

al. (2004) found that after only 3.5 h exposure, significant amounts of toxins were adsorbed

on the resin even though theDinophysis cell numbers were low (100 cells/L). In the

Norwegian trial there was a mixed bloom containing high amounts of different flagellates and

microalgae, typically ca 13 106

cells/L and theD. acuta andD. acuminata numbers ranged

from 100360 cells/L. In this period the HP-20 material also became dark green, indicating a

high concentration of algal pigments in the water. It can not be ruled out that the HP-20

material can become saturated or that the 100 m nylon mesh can clog during the exposure

time and consequently the toxin levels can be underestimated. This needs further

investigation.

3.2 Sample preparation.

Recovery of the lipophilic algal toxins from the HP-20 resin was straight forward. A fresh

water rinse was necessary, prior to elution with MeOH, to remove salts which may disturb

ionization in the HPLC-MS. The ESI interface on the mass spectrometer is susceptible to salt

effects (Gustavsson et al., 2001), and a high salt content can influence the relative intensities

of the H+, NH4

+, and Na

+adducts ions used for MRM quantitation of the toxins. The resulting

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It was found necessary to elute the 3 g of HP-20 resin with at least 2 10 mL of methanol to

fully recover the adsorbed toxins and this is in accordance with the findings of Fux et al.

(2008). It was also important to use a low flow rate through the column, typically 12 bed-

volumes per hour (Manufacturers recommendation). Elution with 23 mL of solvent gave a

diluted sample, but a concentration step can be included depending on the required detection

limits and the sensitivity of the HPLC-MS system. The detection limits obtained with the

instrument used in this work were typically 0.10.3 ng/disk, depending on the toxin. By

omitting the concentration step and adjusting the extract volume to 25 mL, detection limits of

about 25 ng/disk were obtained.

3.3 Toxin profile of disks versus cell counts and blue mussels.

The OA/DTX concentrations in the disks and blue mussels, andDinophysis spp. (D. acuta

andD. acuminata) cell concentrations in the water, are shown in Figure 2. The amount of

OA/DTXs in the disks fluctuated from 120 to 660 ng/g disk, with maxima in weeks 30, 34

and 38. The cell numbers ofD. acuta andD. acuminata also fluctuated during this period,

with numbers ranging from 0 to 360 cells/L and one major peak around week 29. In shellfish,

the sum of both free OA and DTX and their esters was about 65 ng/g at the beginning of the

trial and about 220 ng/g when the trial ended, with peaks at weeks 34 and 40. During the

monitoring period (weeks 2841), three events can be described. The first was the increase of

Dinophysis cell densities and OA/DTX levels in the disks around week 30, but the toxin levels

in the shellfish did not show a corresponding increase (Figure 2). The reason for this might be

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In the second incident, in weeks 3334, levels of OA/DTXs in the disks increased to ca 650

ng/g, and theDinohpysis counts also rose to a moderate level (ca 100 cells/L) in weeks 33 and

34 (Figure 2). The level of OA/DTXs in the shellfish reached a peak in week 34, as did the

levels of OA/DTXs in the disks. OA/DTXs decreased in the shellfish in the following weeks

(3536), when theDinophysis cell numbers and toxin level in the disks also declined.

Depuration of algal toxins from shellfish is poorly understood (Duinker et al., 2007). Passive

samplers could be a useful tool in studies of the depuration of toxins in shellfish through

improved monitoring of the toxin-exposure of the investigated shellfish.

The third event occurred when OA/DTXs in the disks and levels ofDinophysis in the water

reached a maximum in week 38 and 39, respectively (Figure 2). Levels of OA/DTXs in the

shellfish increased in week 39 and reached a maximum in week 40. During this period the

Dinophysis numbers were moderate (ca 140 cells/L) but the amount of other algae was lower

(typically ca 1 106

cells/L) than earlier in the period when the algae population was typically

23 106 cells/L.

Based on these three events, it is difficult to recommend the passive samplers as an early

warning tool. The first incident had increased toxin levels in the disks, with no corresponding

increase in the shellfish. However, in the second and third events there was a marked increase

in OA/DTXs in the disks (weeks 33 and 38) some time before the levels in the shellfish were

observed to increase (weeks 34 and 39).

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week 34 and 38 had 120 (120D. acuminata and 0D. acuta) and 140 (20D. acuminata and

120D. acuta) cells/L, respectively. Algal cell-counting precision is usually good at high cell

densities if over 200 cells of the target species are counted, but poor at low cell densities when

fewer cells are counted (Lund et al., 1958). Except for in weeks 29 and 30, levels of

Dinophysis were below 200 cells/L (corresponding to only 10 cells counted), and this may

account for the lack of a precise correlation between theDinophysis cell counts and the toxin

levels in the disks and shellfish. Lindal, et al 2008 reported substantial variations in toxin

content of bothD. acuta and D. acuminata due to population density and environmental

variations and this may also affect the toxin levels found in the disks and shellfish compared

to the numbers of algae counted.

One difficulty with algal counting as a monitoring tool is that algal blooms can be short-lived

and mobile, and thus occur between algal samplings. This is especially so at locations prone

to tidal flows and/or exposed to wind and wave motion such as in Fldevigen where this trial

was performed. Passive sampling disks should be a valuable tool at such locations, where the

algal counts can change quickly from noDinophysis, up to 360 cells/L and back to a few

cells/L again during one week (Figure 2B). With passive samplers, the water column is

continuously being sampled and hence provides an integrated measurement of toxin levels

throughout the exposure period.

Another problem with algal counting is that it can only provide effective monitoring for toxins

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HP-20 resin has been shown to adsorb OA/DTXs, PTXs, and YTX in New Zealand waters

(MacKenzie et al., 2004) and recently Fux et al. (2008) detected AZAs in Irish waters using

HP-20. In northern European waters, the AZA group is commonly detected in shellfish and is

often present at levels that make shellfish unsuitable for consumption (Hess et al., 2005;

Aasen et al., 2006). During the summer of 2005, blue mussels at Fldevigen contained low

levels of AZA-1, AZA-2, AZA-3 and AZA-6, in a ratio of approximately 3:1:1:0.3

respectively, with concentrations of 2050 g/kg (Figure 4). In the disks, however, only

AZA-1 and AZA-2 (in a ratio of ca 5:1), and no AZA-3 or AZA-6, were detected (Figure 4).

Similarly, Fux et al. (2008) found AZA-1 and AZA-2 in a ratio of ca 4:1 together with traces

of AZA-3, and recently Krock et al. (2008) isolated and cultured an alga producing AZA-1,

AZA-2 and an isomer of AZA-2 but not AZA-3 or AZA-6. This suggests that AZA-3 and

AZA-6 may be produced by metabolism of ingested AZA-1 and AZA-2. Little is known

about the formation and metabolic transformation of the AZAs, but in shellfish a whole range

of AZA analogues has been identified (Satake et al., 1998; Ofuji et al., 1999; Ofuji et al.,

2001; James et al., 2003; Rehmann et al., 2008).

3.5 Detection of SPXs in the disks

A spirolide, most likely 20-methylSPX-G (Aasen et al., 2005), was also detected in the disks

throughout the trial, but only at low levels (ca 540 ng/disk, relative to PTX-2). The MRM

transition of 706.5>164.2, which corresponds to 20-methylSPX-G and SPX-C, was chosen

based on the findings of (Aasen et al., 2005), where 20-methylSPX-G was found to be the

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3.6 PTX-2 and PTX-2 SA in disks.

The detection of both PTX-2 and PTX-2 SA in the disks shows that formation of seco acids

can take place outside of the shellfish, before the algal cells and their PTX-2 are ingested.

PTX-2 SA has previously been observed as a constituent ofDinophysis (Daiguji et al., 1998;

James et al., 1999; Suzuki et al., 2001; MacKenzie et al., 2002)and it appears that conversion

of PTX-2 into PTX-2 SA can be mediated by enzymes present in the algae (MacKenzie et al.,

2002). Esterases responsible for the seco acid formation may leak from damaged algal cells

together with PTX-2, resulting in hydrolysis before adsorption to the HP-20 resin in the disk.

The ratio of PTX-2 to PTX-2 SA in the disks was typically 1:1, compared to 10:1 PTX-2 in

the trial of MacKenzie et al. (2004) in New Zealand, showing that the degree of PTX-2

conversion can vary greatly.

3.7 OA/DTXs and their esters in the disks.

Dinophysis spp. can contain OA diol esters (Suzuki et al., 2004; Miles et al., 2004b; Miles et

al., 2006). However, basic hydrolysis of extracts from passive samplers indicated that little or

no OA or DTX esters were present in the disks. MacKenzie et al. (2004) performed basic

hydrolysis on some of their samples to determine the levels of esterified DTXs, and found

only low amounts of esterified forms (030%) in their extracts. Miles et al. (2004b) isolated a

substantial amount of OA C8-diol ester from harvestedD. acuta cells, and this diol ester was

converted rapidly to OA by a homogenate from the hepatopancreas of the green-lipped mussel

(Perna canaliculus). Also it is known that the complex OA-ester DTX-4 is very short lived

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Concluding remarks.

The passive sampling disk system is cheap to produce and convenient to use and, when

combined with HPLC-MS or ELISA analysis, provides detailed time-averaged information on

the profile of lipophilic toxin analogues in the water. The passive sampling disks have now

been shown to accumulate azaspiracids, okadaic acid analogues, pectenotoxins, yessotoxins

and spirolides. The HP-20 resin in the samplers should also be able to accumulate other

lipophilic algal toxins such as brevetoxin and ciguatoxins. Passive sampling disks have the

potential to be a convenient tool for monitoring the exposure of shellfish and other bivalves to

toxigenic algae containing lipophilic toxins, and may also be useful for monitoring exposure

of aquatic ecosystems to these compounds as well as to a range of lipophilic pollutants.

Acknowledgement

This study was supported by the Norwegian Research Council grant 139593/140, by the

BIOTOX project (partly funded by the European Commission, through the 6th Framework

Programme contract no. 514074, priority Food Quality and Safety, and by the New Zealand

Foundation for Research, Science and Technology (FRST) International Investment

Opportunities Fund (IIOF contract number C10X0406).

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Captions for Figures

Figure 1. Fully assembled passive sampling disk (E), and its component parts: (A) 100 m

nylon mesh; (B) HP-20 resin; (C) inner and (D) outer rings of a 75 mm diameter embroidery

ring with (F) a No. 2 fishing swivel attached.

Figure 2. A) Concentrations of OA/DTXs in passive sampling disks and shellfish (ng/g), and

B)D. acuta +D. acuminata concentration (cells/L) in water for weeks 2841 of 2005.

Figure 3. Typical MRM HPLC-MS chromatogram of toxins in an extract from a passive

sampling disk (week 30) containing 20-methyl-SPX-G, AZA-1, AZA-2, OA, DTX-1, DTX-2,

PTX-2, PTX-12 and YTX.

Figure 4. Chromatogram of AZA profile in extracts of: (A) a passive sampling disk and: (B)

blue mussels (M. edulis).

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A

B

C

D

E

F

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0

100

200

300

400

500

600

700

Concentration

(ng/g) Sum DTX ng/g

disks

Sum DTX ng/g inshellfish

A

0

100

200

300

400

Cellco

unts(cell/L)

28 29 30 31 32 33 34 35 36 37 38 39 40 41

week

BD. acuta + D. acuminata

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0 5 10 15 20

20-methyl-SPX-G

PTX-2SA

OA, DTX-2

PTX-2

AZA-1

AZA-2

DTX-1

YTX

PTX-12a,b

Time (min)

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