Mugil migration studies - MUGIL PROJECT - Université Montpellier 2

Homepage: 

www.mugil.univ-montp2.fr 

Coordinator: 

J. Panfili (IRD) 

DELIVERABLE 4 

MUGIL 

Main Uses of the Grey mullet as Indicator 

of Littoral environmental changes 

2006 - 2009 

February 2008 

Summary 

Deliverable 4 focuses on discussions around 

the migration studies on Mugil cephalus, 

mainly concerning the application of otolith 

chemistry. WS3 has defined guidelines for 

studies on migration using otolith 

microchemistry in order to follow standard 

protocols. 

Mugil migration studies 

European 

Commission 

INCO-CT-2006-026180 


Tomás J., Anastasopoulou K., Berrebi P., Cowley P.D., 

Darnaude A., Diouf P. S., Durand J.-D., Flores-Hernandez D., 

Morales-Nin B., Tzeng W.-N., Wang C.-H., Whitfield A.K., Panfili J. 1,2,3 

1 

Addresses at the end of the document. 

2 

MUGIL Deliverable 4 follows the MUGIL Workshop 3 held in Montpellier (France); 26-28 September 

2007 (see Annexure 1). 

3 

Image on cover: www.fao.org 

INCO-CT-2006-026180 1 MUGIL Deliverable 4 - Feb. 2008

Contents 

1. INTRODUCTION ....................................................................................................................................... 3 

2. USE OF OTOLITHS FOR MUGIL CEPHALUS MIGRATION STUDIES.......................................... 3 

3. OTOLITH PROCESSING FOR THE STUDY OF MUGIL CEPHALUS MIGRATIONS .................. 4 

3.1. FISH COLLECTION AND PRESERVATION ................................................................................................. 4 

3.2. OTOLITH REMOVAL AND PRESERVATION .............................................................................................. 4 

3.3. OTOLITH PREPARATION ........................................................................................................................ 6 

3.3.1. OTOLITH PREPARATION: WHOLE OTOLITH ............................................................................................ 6 

3.3.2. OTOLITH PREPARATION: TRANSVERSE SECTIONS .................................................................................. 7 

3.4. OTOLITH CHEMICAL ANALYSIS ........................................................................................................... 10 

3.4.1. WHOLE OTOLITH ANALYSIS: SB-ICPMS ............................................................................................ 10 

3.4.2. SURFACE ANALYSIS: ELECTRON PROBES WDS ................................................................................... 11 

3.4.3. SURFACE ANALYSIS: LA-ICPMS........................................................................................................ 11 

3.5. DATA TREATMENTS FOR OTOLITH MICROCHEMISTRY ......................................................................... 11 

4. GENERAL CONCLUSIONS AND RECOMMENDATIONS.............................................................. 13 

5. REFERENCES .......................................................................................................................................... 13 

6. AUTHOR ADDRESSES ........................................................................................................................... 15 

7. ANNEXURE 1 – WS3 AGENDA ............................................................................................................. 16 

INCO-CT-2006-026180 2 MUGIL Deliverable 4 - Feb. 2008 

Page

1. Introduction 

Mugil cephalus is a diadromous fish species that migrates between continental and marine 

environments during its life cycle: juveniles and sub-adults grow in freshwater and brackish water 

habitats (estuaries and lagoons). Adults undertake off-shore migrations to sea for spawning, usually in 

the form of large schools (Thomson, 1955; Bacheler et al., 2005). 

The aim of this deliverable is to define the best method for otolith collection, storage, preparation 

and microchemical analysis to study the transhaline migrations of grey mullets. The guidelines 

presented here will enable all future researchers to follow the same standard protocols for migration 

studies involving otolith microchemistry in M. cephalus, thereby allowing global comparisons on this 

subject. 

2. Use of otoliths for Mugil cephalus migration studies 

Otolith microchemistry has proved successful in resolving habitat occupation between freshwater 

and seawater environments by diadromous fish, such as salmonids (Kalish, 1990; Volk et al., 2000; 

Arai et al., 2007), eels (Jessop et al., 2002; Tzeng et al., 2003; Daverat et al., 2005) and shads 

(Thorrold et al., 1998; Tomás et al., 2005). Otolith microchemistry has also been successfully applied 

to the identification of saline habitats occupied by flathead (grey) mullet Mugil cephalus but these 

studies are scarce and almost completely confined to Taiwan (Chang et al., 2004a; Chang et al., 

2004b). While catadromy was supposed to be obligatory for M. cephalus these studies have shown 

that it is only a potential strategy for newly recruited fry. Indeed, otolith microchemistry has shown 

that some individuals avoided entering continental waters and continued moving between estuaries 

and the sea. In the absence of detailed studies on saline habitat use by M. cephalus from other parts of 

the world, it already appears that otolith microchemistry constitutes a very efficient approach to 

identify and determine the regional range of migration of the species. 

Otolith microchemistry has also brought new evidences regarding nursery fidelity by M. cephalus, 

again in the waters of Taiwan. In an attempt to study fidelity to nursery areas by M. cephalus, otolith 

trace elemental composition was used as an indicator to discriminate the juveniles from various 

estuaries of Taiwan (Wang et al., 2006). Out of the twelve elements (Li, Na, Mg, K, Mn, Fe, Ni, Cu, 

Zn, Sr, Ba, and Pb) analysed from the otoliths of juvenile mullet, five elements (Mn, Ni, Zn, Sr, and 

Ba) were found to be significantly different between estuaries. The canonical discriminant analysis 

also indicated that 10 of 12 elemental ratios (except Li/Ca and Cu/Ca) played a significant role in the 

discrimination of juvenile M. cephalus among the estuaries. Among the elements, Mn/Ca, Ni/Ca and 

Zn/Ca contributed approximately 49.8 % in the first canonical function of the discrimination, while 

Ba/Ca and Sr/Ca contributed 32.9 % in the second function. As a result, 84 % of the early juveniles 

could be assigned to their recruited estuaries with their otolith chemical signatures, indicating that 

elemental composition in the otolith can be used as a natural tag to trace their nursery areas (Wang et 

al., 2006). 

While the use of otolith microchemistry seems promising for the study of M. cephalus migrations, 

several constraints exist in the use of this technique. Otoliths are often purer carbonates than other 

calcified structures (Campana, 1999) and the elemental signatures that are used in otoliths to 

discriminate between water streams, coastal waters, etc. are in the order of ppm or ppb. Thus, it is 

critically important to avoid otolith contamination by external elements (often extremely ubiquitous) 

during otolith extraction, manipulation and preparation. Moreover, the high number of analytical tools 

used for these studies are also sources of differences in the results (Campana et al., 1997). 

The purpose of MUGIL WS3 was to identify the critical steps in otolith processing which could 

lead to alterations in otolith composition and minimise the sources of contamination at each step. The 

usefulness of several otolith analysis techniques for describing M. cephalus migrations was also 

compared and the most valuable analytical instruments were identified and listed by the MUGIL 


consortium. Therefore, WS3 aimed to develop a standardised protocol and method for the application 

of otolith microchemistry to the study of M. cephalus migrations. 

3. Otolith processing for the study of Mugil cephalus 

migrations 

Foreword: During otolith processing, contact between the otolith surface and metallic tools, bare 

hands or any liquid of dubious composition or low purity should be avoided. 

3.1. Fish collection and preservation 

Fish should preferably be manipulated and dissected just after their capture. If this is not possible fish 

should be frozen immediately after capture to prevent ionic exchanges between the plasma and the 

endolymph (internal fluid that fills the otosac and is in direct contact with the otolith). If immediate 

freezing is not possible, the fish can be kept refrigerated for a few hours before otolith removal (within 

the day). The use of formaldehyde or alcohol (even trace metal grade 100% ethanol) to preserve fish 

or fish heads must be avoided since it can alter otolith composition. The following procedure must be 

followed for otolith extraction in the laboratory (otolith dissection in the field is described at the end of 

this section). 

3.2. Otolith removal and preservation 

Disposable (powder free) polyethylene gloves must be worn at all times during otolith removal 

and subsequent handling. All the material to be in contact with either the otoliths or the solutions used 

to clean them (i.e. all dissecting tools and storage equipment) must be plastic, preferably Teflon, 

otherwise in high-density Polyethylene (HDPE) and need to be decontaminated prior to use. 

a) Procedure for material decontamination prior to use 

This procedure involves the use of a "Clean Cell" (i.e. class 10-horizontal laminar flow hood with 

its floor lined with polyethylene sheeting replaced periodically and wiped off before and after use with 

ultra-pure trace-metal grade ethanol). Only the material which has been decontaminated is referred to 

as "clean" in the present document. The decontamination procedure is as follows: 

1) Wash all equipment in 95% non-denatured ethanol. 

2) Rinse in ultra-pure (trace metal grade, 18.2 MΩ) water. 

3) Rest-wash (≥ 24 hours) in 4% nitric acid within HDPE buckets covered before and during 

immersion (acid bath = 0.2 L of 60% trace-metal grade HNO3 diluted with 2.8 L of ultra-pure 

water). 

4) Rinse 3 times with ultra-pure water. 

5) Rest-rinse (≥ 24 hours) in ultra-pure water within clean HDPE buckets covered before and 

during immersion. 

6) Air-dry for 12 h within the Clean Cell. 

7) Repackage in sealed clean plastic bags. 


) Procedure for otolith extraction and preservation in the laboratory 

Use clean plastic (Teflon or HDPE) tools only to handle the otoliths and avoid metallic tools as 

much as possible during fish dissection. Wear gloves on both hands at all times (to manipulate the 

fish but also the dissecting instruments). All tools must be cleaned both before use and between 

each new dissection with 10% trace-metal grade nitric acid, then ultra-pure water. If the fish is 

frozen, it should be defrosted prior to otolith extraction. The fish dissection and otolith extraction 

protocol is as follows: 

1) Put an extra glove on the hand holding the fish, which will be removed after otolith extraction 

(i.e. when disposing the dissected fish). 

2) Hold the fish and make a frontal incision right above its eyes with a knife. For more detail 

regarding this section, refer to the Manual of Fish Sclerochronology (Panfili, 2002). NB: avoid 

scratching the otolith with the knife during this step. 

3) Using plastic forceps, crack the head open and remove fish’s brain. 

4) Once the otoliths are visible, use a second (clean) pair of plastic forceps to extract the otoliths. 

5) Place each otolith in a clean Petri dish filled with ultra-pure water (keep the Petri dish closed 

before use). 

6) Close the Petri dish, discard the fish and remove the extra glove on the hand holding the fish. 

7) Use the closed Petri dish for transporting the otoliths to view under a stereomicroscope. 

8) Using reflected light, clear the surface of each otolith of any adhering organic tissue using 

plastic forceps. 

9) Immerse the otolith for five minutes in a new clean microtube vial filled with 20% hydrogen 

peroxide (H2O2) (keep closed) to oxidise all remaining organic material from the otolith 

surface. 

10) Rinse the otolith in a new clean microtube vial filled with ultrapure water (keep closed). 

11) Once rinsed, place each otolith in a clean perfectly dry uniquely labelled plastic microtube 

vial (kept closed until this step). 

12) Close the microtube vial and reference it (fish identification number and indication of 

whether it is a LEFT or a RIGHT otolith). Left and right otoliths should be stored separately in 

different microtube vials since one could be used for microchemistry, and the other for age 

estimation. Information recorded for each referenced otolith should include the following: (i) 

unique reference number (engraved on the microtube vial), (ii) species, fish lengths (standard, 

fork and/or total) and mass, (iii) collection location (including GPS co-ordinates), (iv) 

collection method, and (v) collection date. 

13) Under the Clean Cell, open the microtube vials leaving the otoliths inside and allow drying for 

24 h under the ultra-clean conditions. 

14) Close the microtube vials containing individual otoliths and store them in the laboratory in 

sealed plastic bags until otolith processing. It is recommended to store only the otoliths from 

the same sampling station or sampling date in the same sealed plastic bags. Microtube vials 

containing LEFT and RIGHT otoliths should be stored in separate plastic bags to avoid 

unnecessary manipulation of vials since LEFT and RIGHT otoliths could have different uses. 

c) Notes on otolith extraction and preservation in field conditions 

Discussions during WS3 identified a number of cases where the above-mentioned ideal procedures 

(i.e. under clean laboratory conditions) would be impractical. These include: (i) when large numbers 

of fish are captured at a time, (ii) when dealing with exceptionally large fish, or (iii) when field trips 

last for more than a day and fish storage (freezing) capacity is limited. Under such conditions the 

following procedure is recommended from step 7: 

7) Place otoliths in a Petri dish and grossly clean the otolith surface of any adhering organic 

tissue, using plastic forceps. 


8) Eventually sonify the otoliths in clean vials in ultrapure water during 3 to 5 minutes. 

9) Place each washed otolith in a clean uniquely labelled plastic microtube vial (perfectly dry 

and kept closed until this step). 

10) Close the microtube vial and reference it (fish identification number and indication of 

whether it is a LEFT or a RIGHT otolith). Left and right otoliths should be stored separately in 

different microtube vials since one can be used for microchemistry, and the other for age 

estimation. 

NB1: the otoliths will only be H2O2 cleaned and left to dry out (steps 9 to 14 of the lab procedure) 

once back at the laboratory. 

NB2: the otolith is a porous calcium carbonate structure. Its composition can be altered by 

incorporation of contaminating elements in fluids where the otolith is immersed, or tools with which 

the otolith surface is in contact, but also by leaking out of elements originally present in the otolith. 

For this reason it is important to avoid unnecessary manipulation of the otolith and extreme care 

during otolith manipulation should practised in these initial stages. 

3.3. Otolith preparation 

Otolith preparation is a required step to extract chemical information. Relevant microchemistry 

information can be acquired EITHER by analysing each otolith as a whole (thus obtaining a unique 

chemical signature for each individual corresponding to its complete life-time) OR by sampling 

discrete areas between age marks within the otolith (typically 5 to 50 μm-wide spots made on otolith 

transverse sections) that can be related to particular periods of the life of the fish. Before analysis, an 

image of the whole and/or sectioned otolith should be recorded but clearing media should be avoided 

(the only media should eventually be ultra-pure water). 

3.3.1. Otolith preparation: whole otolith 

The analysis of whole dissolved otoliths is the easiest, quickest and least contaminant way to 

extract chemical information from the otoliths. Unfortunately, this approach does not allow linking 

microchemistry variation to otolith age marks, unless the analysed individuals belong to the same age 

class and are captured at the same time. Therefore, the analysis should mainly be used to assess 

nursery area fingerprints from juvenile fish from different origins. 

The analysis of whole otoliths should be carried out individually. The first steps of this analysis 

should follow the protocol below: 

1) Weigh the otoliths individually with a 0.001 mg precision. Otolith weight needs to be 

accurately measured as elemental concentration is reported on a weight basis. Otolith 

weighing and manipulation should be carried out in ultra-clean conditions using dry plastic or 

Teflon forceps (previously cleaned with 10% nitric acid). 

2) Once weighed, each otolith should be transferred to individual plastic vials waiting for acid 

dissolution. 

3) Acid dissolution of the otolith should be carried out with ultra-pure nitric acid using a 

determined volume. Once dissolved the otolith solute is ready for analysis (See Section 3.4 on 

Otolith Chemical Analysis). 


3.3.2. Otolith preparation: transverse sections 

Otolith transverse sections have already been identified during MUGIL WS2 as the most 

appropriate way of exposing internal growth marks in the otolith for age estimation. As such, chemical 

information to be related to these marks will be extracted from transverse sections. 

While detailed information on how to prepare otolith transverse sections for age estimation is 

available in MUGIL Deliverable 3 (Mugil life-history trait studies, October 2007), the preparation of 

otolith sections for microchemistry analysis requires additional precautions to avoid alteration of the 

original composition of the otolith. As for otolith dissection, the contact of the otolith with metallic 

tools should be avoided. 

a. Embedding 

The dried otolith is embedded in a transparent resin but polyester resins, as used for otolith 

preparation for age estimation, should be avoided as their composition might be a source of 

contaminating elements. Epoxy resins are preferred, among which Araldite seems a good compromise. 

The use of individually labelled moulds (e.g. made of silicon elastomer), dimensioned according to the 

otolith size is necessary. The below protocol should be followed: 

1) The resin and its catalyst are mixed thoroughly but taking care to prevent bubbles forming in 

the resin mixture. The process should only involve the use of plastic tools (vials, dishes, 

pipettes, even the stick used to mix the resin and the catalyst). 

2) The mixture is then left to rest for a few minutes to allow the biggest bubbles to reach the 

surface and disappear. 

3) The polyester resin is pipetted into the mould cell to create a base layer and allowed to harden 

for 24 h in an ultra clean dry oven (around 30-35 °C). 

4) The otolith is then placed in a mould cell and new resin (preparation as in 1)) is poured over 

the otolith until it is completely covered. At this step it is important to turn over the otolith in 

its mould to eliminate air bubbles that are often trapped below. After this, the otolith needs to 

be re-positioned before resin hardens. 

5) Re-positioning takes into account the next steps of preparation (e.g. the sectioning plane): the 

otolith is embedded with it anterior-posterior axis parallel to the main axis of the mould. 

6) The preparation is then set to harden for 24-48 h in a dry oven (30-35°C). The embedded 

otolith is then ready for sectioning. 

Note that any manipulation of the otolith, from the plastic vial where it is stored to the mould, 

even when immersed in the resin before it hardens, has to be done with plastic or Teflon forceps. The 

resulting resin block should not be manipulated with bare hands, always use clean gloves to remove it 

from the mould. If the block is not to be cut right after embedding, it should be stored in sealed plastic 

bags in the same manner as the microtube vials containing freshly extracted otoliths. 

b. Transverse sectioning 

The sectioning reveals the internal otolith increments. While being a necessary step it also 

represents a critical stage at which the internal structure of the otolith is exposed to fluids and 

suspensions. Extra care has to be taken to follow a standard procedure. Sectioning is done using a lowspeed 

circular saw (e.g. Isomet® saw, Buehler Ltd) equipped with a diamond blade. The cooling fluid 

used has to be ultrapure MilliQ water. Usually the recipient used for the cooling liquid is made of bare 

metal. While one possibility is to find a container of similar dimensions made of plastic, it is also 

possible to place a smaller container made of plastic inside the main metallic container. This avoids 

any contact between the cooling fluid and the metal and thus between the blade and subsequently the 

otolith internal structure. As in the previous stage, always manipulate the resin block using clean 

gloves. It is not possible to replace the metallic arm of the saw, the metallic resin block holder or the 

metallic instruments to tighten the block to the holder by non metallic substitutes. Thus care has to be 


taken to avoid unnecessary friction between these and the resin block. The next steps of the procedure 

should follow the protocol below: 

1) Fill the water container of the saw with ultrapure MilliQ water. 

2) Fix the resin block including the otolith to the saw holder. After this step no further 

manipulation of the block is required. 

3) Adjust the section levels with the saw micrometer. Work to obtain sections with a maximum 

500 μm if the otolith is to be sampled by Laser Ablation and a maximum of 300 μm if the 

otolith surface is to be analysed with an electron probe (See Section 3.3 on Chemical 

Analysis). 

4) Align the diamond blade at the level of the otolith core. 

5) Move the diamond blade with the saw micrometer at a distance of half of the required section 

thickness. Section the resin block. 

6) Move the diamond blade with the saw micrometer in the opposite direction at a distance of the 

required section thickness. Section the resin block. The resulting section will be detached from 

the resin block at the end of the second section. Avoid having the section falling down into the 

recipient containing the cooling fluid. Hold it with thin plastic or Teflon forceps and remove it 

from the sectioning set to a Petri dish to be rinsed with MilliQ water. 

7) Place inside the section into a microtube vial and leave to dry. 

8) Return to the sectioning set, remove the resin block and thoroughly rinse the blade with 

ultrapure MilliQ water. 

9) Dispose the water from the saw container, rinse thoroughly and fill again with fresh ultrapure 

MilliQ water, ready for the next otolith. 

c. Section manipulation 

If the dry otolith section cannot be further manipulated without being affixed to a solid support, 

due for example to its small size, the section should be attached to a clean glass slide (preferably of the 

geological type rather than the histological type). Use Araldite resin to mount the section onto the 

centre of the glass slide and manipulate the section with plastic or Teflon forceps. The glass slide 

should not be manipulated without clean gloves. 

d. Surface polishing 

Growth marks will be best observed in a plane that includes the core. Since the objective of 

sectioning is only to grossly expose the internal structure of the otolith, grinding and polishing are 

necessary to reach the plane of the core. Some degree of experience is necessary to make thin sections 

of otoliths. Otolith preparation for microchemistry analysis should be attempted after the user is 

familiarised with the preparation of thin sections for age estimation. 

Grinding should use clean abrasive papers wetted with ultrapure MilliQ water only. In all cases 

the use of an automatic polishing machine should be regarded as highly desirable since it will ensure a 

standard grinding and polishing of preparations. 

1- Use gloves to manipulate abrasive papers between the site of storage (this must be a dust free 

holder in closed cupboards) and the polishing machine. 

2- Place the abrasive paper in the polishing machine. 

3- Attach the otolith section mounted on a geological glass slide to the specimen holder of the 

polishing machine. 

4- Place the specimen holder on the abrasive paper and under the arm of the polishing machine. 

5- Wet the abrasive paper with ultrapure MilliQ water. 

6- Start the grinding process. 

7- Check regularly the level of grinding under a binocular microscope. To do so, it will be 

necessary to slightly polish the otolith surface to remove scratches left by the grinding. To do 

so, proceed as follows: 

8- Remove the specimen holder with the otolith from the polishing machine. 

9- Clean in an ultrasonic bath with ultrapure MilliQ water. 


10- Rinse the holder with the otolith with ultrapure MilliQ water. 

11- Polish the otolith against a polishing cloth with a 3 microns water based diamond suspension 

(the manipulation of polishing cloths are the same as for abrasive papers, as described in 1)). 


13- Place the specimen holder with the otolith under a binocular microscope and check the level 

of polishing. If the plane of the core is not near the otolith surface, move to steps (9), (10) and 

from (2) to (13) until reaching the plane of the core. 

14- Once finished, remove the glass slide with the otolith from the specimen holder. 

15- Clean it in an ultrasonic bath with ultrapure MilliQ water during 1-3 minutes. 

16- Place it in a clean plastic slide box (kept closed until then to avoid dust) and leave to dry 

under the Clean Cell and allow drying for 24 h under the ultra-clean conditions. 

17- Clean the specimen holder of the polishing machine with ultrapure MilliQ water. 

18- Rinse thoroughly the abrasive paper of the polishing machine with ultrapure MilliQ water. To 

insure less contamination the abrasive paper should be changed between otoliths. 

19- Proceed to the grinding of the next otolith. 

If samples are to be analysed with electron probes the surface of the otolith section needs to be 

perfectly flat and free of scratches. Thus, final polishing has to be taken to the finest grains of 

polishing (1 μm to 0.25 μm) to ensure the removal of scratches. Always check the quality of the 

preparation under an epi-illumination compound microscope at high magnification. If samples are to 

be analysed with LA-ICPMS, the quality of the preparation is less relevant and a polishing with 3 to 

1 μm should be enough. In all cases, polishing suspensions should be water based diamond 

suspensions (alumina powders should be avoided). For polishing, proceed as follows: 

20- Use gloves to manipulate the polishing cloth between the site of storage (this must be a dust 

free holder kept in closed cupboards) and the polishing machine. 

21- Place the polishing cloth in the polishing machine. 

22- Attach the otolith section mounted on a geological glass slide to the specimen holder of the 

polishing machine. 

23- Place the specimen holder on the polishing cloth and under the arm of the polishing machine. 

24- Wet the polishing cloth with a 3 µm water based diamond suspension. 

25- Start the polishing process. 

26- Check regularly the progress of the polishing process under a compound microscope with epiillumination. 

At this stage, it is a matter of assessing whether scratches disappear from the 

otolith surface. To do so, proceed as follows: 

27- Remove the specimen holder with the otolith from the polishing machine. 

28- Clean in an ultrasonic bath with ultrapure MilliQ water during few seconds. 


30- Place the glass slide with the otolith under a compound microscope with epi-illumination at 

×50 magnification and check the progress of the polishing. If more polishing with the 3 µm 

suspension is required, move to steps (28), (29) and from (23) to (26) until the quality of the 

surface is satisfactory across the surface of the sample. 

31- Once finished, remove the glass slide with the otolith from the specimen holder. 

32- Clean it in an ultrasonic bath with ultrapure MilliQ water during 1-3 minutes. 

33- Place it in clean plastic slide box and leave to dry under the Clean Cell and allow drying for 

24 h under the ultra-clean conditions. 

34- Clean the specimen holder of the polishing machine with ultrapure MilliQ water. Change the 

polishing cloth to avoid contamination. 

35- Proceed to the polishing of the next otolith. 

The same sequence of steps from (20) to (35) will be followed using 1 µm water diamond 

suspensions. An additional sequence of polishing with a 0.25 µm water diamond suspension will be 

done if the otolith surface is to be analysed with WDS. 


e. Final cleaning 

After the polishing stage, the preparation should be cleaned in an ultrasonic bath with MilliQ 

water during 1-3 minutes. Place it in a clean plastic slide box (kept closed until then to avoid dust) and 

leave to dry under the Clean Cell for 24 h under the ultra-clean conditions. 

f. Image acquisition 

The otolith increments on the transverse sections are to be examined under transmitted light. It is 

recommended that digital images of the otolith sections are captured showing relevant parts of the 

otolith, i.e. core, growth marks along the axis of analysis and otolith edge as these will ease locating 

the zone of the otolith to be analysed once the otolith is inside the analytical instrument. 

3.4. Otolith chemical analysis 

The chemical analysis of the otolith seeks extracting information on elements or isotopes through 

screening and quantification of concentrations from the otolith mineral and organic matrix. It is 

possible to extract information from the whole otolith by sampling the solution resulting from 

dissolving the whole otolith OR by sampling the otolith at discrete spots using high resolution electron 

probes OR using laser ablation which is the more powerful technique. The choice of instruments to be 

used within MUGIL was decided during WS3 and selected instruments were chosen among several 

candidates listed in the literature (EDS, ICPMS, SIMS, AAS, AES…). All these approaches have 

advantages and disadvantages which are outlined below. 

3.4.1. Whole otolith analysis: SB-ICPMS 

Dissolving and quantifying elements in the whole otolith will provide information about the 

concentrations of elements over the entire life of the fish. Since this approach does not require more 

preparation than dissolving the otolith in nitric acid, otolith composition is probably maintained. Yet, 

this approach will render data of limited use to study fish migration: while it would probably help 

discriminating migrating fish from non-migrating fish, it will not help to distinguish a fish that 

commutes every season from seawater to freshwater from a fish that has spent half of its life in 

freshwater and the other half at sea. Yet, it can be regarded as an interesting approach to make 

estimations about the fraction of a sample/population that is freshwater resident, seawater resident or 

migrant. 

The selected instrument to be used within MUGIL is an Inductively Coupled Plasma Mass 

Spectrometer (ICPMS) sampling in continuous mode the solution containing the dissolved otolith. The 

solution is pumped towards the plasma torch which ionises the sampled matrix. Ionised elements are 

accelerated through cones to align and sub-sample the ionised cloud. Magnets and mass acceleration 

serve to filter masses of elements which are sequentially projected to a Faraday cup or electron 

multiplier for quantification. Counts are then related to concentrations by use of standards analysed at 

increasing concentrations. Internal standards should be quantified over any session of analysis to 

quantify instrument drift. Samples should be analysed in random sequences. Interferences between 

molecules of same mass and particularly those containing the ubiquitous oxygen or calcium should be 

checked (instruments these days include lists of interfering signals and have also increased their mass 

resolution). 

The advantages of this instrument are i) the low limits of detection that allow quantifying isotopes 

at the ppb level (part per billion), ii) the fact that this type of instrument is increasingly common in 


geological labs and universities, and iii) the possibility of processing a large number of samples for 

which a large number of elements or isotopes can be quantified at the same time. 

Fish migration will definitely be best studied by analysing the variation of concentrations of the 

elements selected as tracers of salinity changes (usually Sr and Ca) in relation to age marks in the 

otolith. This implies sampling only the surface of the otolith at discrete spots. The required size of 

these spots, the number and expected concentrations of the elements to be measured and the cost of 

analysis will determine the use of an electron probe (WDS) or a laser ablation ICPMS instrument. 

3.4.2. Surface analysis: electron probes WDS 

Low costs and easy-access are the main reasons to choose an electron probe to analyse Sr and Ca 

in fish otoliths. The selected type of electron probe to be used within MUGIL is a Wavelength 

Dispersive Spectrometer (WDS) which has the advantage of separating the functions of detection and 

quantification as opposed to other types of electron probes. Analysis is based on the detection and 

quantification of X-Rays emitted from a discrete zone of the otolith surface excited by a beam of 

primary electrons accelerated from a tungsten filament located above the sample. To improve the 

quality of analysis the otolith surface should be previously coated with a thin layer of carbon to 

remove electrons accelerated to the sample. Beam current, accelerating voltage and spot size should be 

standardised to 10 nA, 15 keV and 10 μm within MUGIL consortium. Counting times at peak should 

be 30 sec for Ca and 90 sec for Sr and half in the background (only one side). 

3.4.3. Surface analysis: LA-ICPMS 

Otoliths will be best screened for elements with Laser Ablation Inductively Coupled Mass 

Spectrometry (LA-ICPMS) when the elements to quantify are more than Ca and Sr. For instance, 

alternative elements such as Ba may constitute good tracers of estuarine environments. To do this, the 

otolith section is introduced in the sealed laser chamber coupled to an ICPMS instrument. A CCD 

camera inside the chamber allows visualisation of the otolith surface and selecting the exact location 

to be hit by the laser from a monitor. Firing the laser at the otolith surface will result in the ablation of 

otolith portions that will be left in suspension over the otolith. An Argon gas flow transports any 

vaporised matter ablated towards the plasma torch within the ICPMS where the ionised elements are 

filtered, aligned and quantified (See Section 3.4.1.Whole otolith analysis: SB-ICPMS). Sizes of the 

ablated spots vary between 50 down to 5 μm. Any element present in these vaporised portions of the 

otolith surface whose concentration which is above the limit of detection of the instrument can be 

measured in the same manner described above in Section 3.4.1. (Whole otolith analysis: SB-ICPMS) 

taking advantage of the capabilities of the ICPMS. 

3.5. Data treatments for otolith microchemistry 

The extraction of data of chemical composition through surface analysis pursues the 

characterisation of elemental variations across the otolith, usually between the otolith core (formed at 

hatching) and the otolith edge (formed just before capture). 


The standards to be used by each laboratory should be the same. If different glass standards are 

used with LA-ICPMS instruments, an inter-calibration exercise should be run before running 

extensive analyses between the laboratories implicated. 

The typical output of data is presented in Table 1 and can be imported as a spreadsheet where lines 

represent the information relative to each isotope (Ca 44 , Sr 88 for example) and columns list each 

sample, standards or blanks. 

Data in CPS (counts per second) are transformed through the calibration curves from the standards 

into concentrations in ppm (parts per million) or ppb (parts per billion). Depending on the number of 

samples to be treated, it may be necessary to use internal standards to assess for instrumental drift 

along the day, for example by alternating the analysis of otoliths and standards. Raw data should be 

inspected first to assess instrument drift. 

Raw data have to be treated before being used. To do so, follow the following protocol: 

1. Identify in the columns which are standards, internal standards and samples. 

2. Look for raw data with background subtracted (clean raw data). 

3. Look for MDL (Minimum Detection Limits) data that represent the threshold below which 

the instrument cannot distinguish between concentrations and noise. Note that these are 

generally reported in ppm or ppb. 

4. Look for clean raw data in the ppm or ppb form (not CPS) not filtered for MDL. 

5. Verify that for each analysis and isotope, concentrations are above the MDL. The instrument 

software may do this automatically. Any data below MDL should be discarded. The resulting 

data is MDL filtered and can be used for posterior data treatment. 

Table 1. LA-ICPMS data output of chemical composition: isotopes of calcium (Ca), strontium (Sr) and barium (Ba). 

GLITTER4.0: Laser Ablation Analysis Results 

E:\LA-DATA\JacquesPanfili\mmh403profil\MMH403profil.FIN 

Mon Jan 21 10:21:07 2008 

All values are reported in ppm 

GLITTER!: Trace Element Concentrations MDL filtered. 

Element DCSTD_11 MMH403profil_1 MMH403profil_2MMH403profil_3 MMH403profil_4 MMH403profil_5 DCSTD_12 

Ca43 81833,28 400232,66 400232,59 400232,63 400232,59 400232,63 81833,28 

Ca44 82162,27 406706,06 411860,66 408979,72 409102 410456,66 81807,77 

Sr88 499,63 3447,93 2042,35 1515,89 2321,84 2202,8 496,31 

Ba138 425,18 6,89 2,904 5,65 13,19 12,75 423,41 

GLITTER!: Trac Not filtered for MDL. 


Ca43 81833,28 400232,66 400232,59 400232,63 400232,59 400232,63 81833,28 

Ca44 82162,27 406706,06 411860,66 408979,72 409102 410456,66 81807,77 

Sr88 499,63 3447,93 2042,35 1515,89 2321,84 2202,8 496,31 

Ba138 425,18 6,89 2,904 5,65 13,19 12,75 423,41 

GLITTER!: 1 sigma error. 


Ca43 2630,77 12692,09 12690,12 12689,5 12689,15 12692,33 2629,86 

Ca44 2620,54 12876,02 13038,11 12946,58 12950,26 12994,88 2608,74 

Sr88 15,34 105,02 62,21 46,18 70,72 67,1 15,24 

Ba138 12,97 0,22 0,1 0,18 0,41 0,4 12,92 

GLITTER!: Minimum detection limits (99% confidence). 


Ca43 282,65 231,63 219,19 226,68 213,67 219,31 256,4 

Ca44 101,55 85,52 81,24 83,98 79,76 84,45 95,11 

Sr88 0,178 0,143 0,148 0,148 0,149 0,155 0,173 

Ba138 0,0616 0,05 0,048 0,05 0,0448 0,0456 0,0559 

GLITTER!: Mean Raw CPS background NOT subtracted. 


Ca43 179858 958731 1013360 1007298 1042919 1002256 184573 

Ca44 2830746 14773582 15805902 15605130 16160239 15591435 2894563 

Sr88 1042617 8113390 5084012 3751397 5948820 5425059 1068471 

Ba138 976801 18507 8601 16066 37884 35195 1003332 

GLITTER!: Mean Raw CPS background subtracted. 


Ca43 171853 951553 1006173 1000009 1035684 995495 177271 

Ca44 2596698 14551988 15582310 15378525 15931904 15364383 2667015 

Sr88 1037971 8109365 5079231 3746864 5943681 5420118 1063571 

Ba138 976119 17908 7982 15429 37313 34671 1002709 


4. General conclusions and recommendations 

Following MUGIL WS3 the consortium has defined guidelines for the application of otolith 

microchemistry to study migration and dispersal across saline gradients by Mugil cephalus and 

proposes a standard protocol for otolith dissection, manipulation, storage, preparation and analysis. 

For transhaline migration studies of M. cephalus the MUGIL consortium recommends the 

chemical analysis of transverse sections of otoliths. Sections should be preferentially analysed with 

LA-ICPMS along the axis of the otolith with visible and interpretable growth marks. 

Extreme care should be put in manipulating otoliths and avoiding excessive manipulation, the 

contact with any metallic tool, bare hands or immersion or rinsing with solutions of unknown 

composition and low purity. As such, otoliths should only be rinsed in ultrapure MilliQ water and 

cleaned in pure hydrogen peroxide (H2O2). Otoliths can only be stored dry and clean of any organic 

remains in individual microtube vials in clean sealed plastic bags. While it is acknowledged that it is 

not feasible to avoid altering to some extent the composition of the otolith by manipulation and 

preparation, it is stressed that the strict use of this standard protocol will reduce the sources of error. 

Otolith analysis should be done with LA-ICPMS (for multi-elemental screening) or WDS (for Sr 

and Ca). It is acknowledged that Sr and Ca are to date the best candidates to track transhaline 

migrations of individual fish, yet the possibility exists that other elements may bring additional 

information to identify fish dispersal within freshwater using strontium isotopes or to identify fidelity 

to certain coastal areas using Mn, Ni, Zn and Ba. Since LA-ICPMS is superior to WDS in multielemental 

screening and sensibility, the former should be used with otoliths. 

Otolith microchemistry has drawn much attention in recent years and the number of publications 

reporting results on the application of otolith microchemistry to study diadromous fish transhaline 

migrations has increased significantly. Yet more and more evidence is available regarding the 

necessity of validating the relationship between otolith chemical information and environmental 

information. Therefore, the MUGIL consortium recommends the conducting of validation experiments 

in controlled rearing conditions before applying the technique. Experimental designs will bring the 

possibility of working out coefficients of discrimination of relevant elements between the water and 

the otolith for fish of different ages/sizes in at least two zones covered by MUGIL. Validation 

experiments will allow for a wide application of the technique based on a sound methodology. 

5. References 

Arai, T., Hirata, T., & Takagi, Y. (2007). Application of laser ablation ICPMS to trace the 

environmental history of chum salmon Oncorhynchus keta. Marine Environmental Research 

63 (1), 55-66. 

Bacheler, N. M., Wong, R. A., & Buckel, J. A. (2005). Movements and mortality rates of stripped 

mullet in North Carolina. North American Journal of Fisheries Management 25 (1), 361-373. 

Campana, S. E. (1999). Chemistry and composition of fish otoliths: pathways, mechanisms and 

applications. Marine Ecology Progress Series 188, 263-297. 


Campana, S. E., Thorrold, S. R., Jones, C. M., Gunther, D., Tubrett, M., Longerich, H., Jackson, S., 

Halden, N. M., Kalish, J. M., Piccoli, P., de Pontual, H., Troadec, H., Panfili, J., Secor, D. H., 

Severin, K. P., Sie, S. H., Thresher, R., Teesdale, W. J., & Campbell, J. L. (1997). 

Comparison of accuracy, precision and sensitivity in elemental assays of fish otoliths using the 

electron microprobe, proton-induced X-ray emission, and laser ablation inductively coupled 

plasma mass spectrometry. Canadian Journal of Fisheries and Aquatic Sciences 54, 2068- 

2079. 

Chang, C. W., Iizuka, Y., & Tzeng, W. N. (2004a). Migratory environmental history of the grey 

mullet Mugil cephalus as revealed by otolith Sr:Ca ratios. Marine Ecology Progress Series 

267, 277-288. 

Chang, C. W., Lin, S. H., Iizuka, Y., & Tzeng, W. N. (2004b). Relationship between Sr:Ca ratios in 

otoliths of grey mullet Mugil cephalus and ambient salinity: validation, mechanisms, and 

applications. Zoological Studies 43 (1), 74-85. 

Daverat, F., Tomás, J., Lahaye, M., Palmer, M., & Elie, P. (2005). Tracking continental habitat shifts 

of eels using otolith Sr/Ca ratios: validation and application to the coastal, estuarine and 

riverine eels of the Gironde-Garonne-Dordogne watershed. Marine and Freshwater Research 

56, 619-627. 

Jessop, B. M., Shiao, J. C., Iizuka, Y., & Tzeng, W. N. (2002). Migratory behaviour and habitat use by 

American eels Anguilla rostrata as revealed by otolith microchemistry. Marine Ecology 

Progress Series 233, 217-229. 

Kalish, J. M. (1990). Use of otolith microchemistry to distinguish the progeny of sympatric 

anadromous and non-anadromous salmonids. Fishery Bulletin 88 (4), 657-666. 

Panfili, J. (2002). Extraction and conservation of calcified structures. In Manual of Fish 

Sclerochronology (Panfili, J., de Pontual, H., Troadec, H., and Wright, P. J., eds.), pp. 317- 

330. Brest, France: Ifremer-IRD coedition. 

Thomson, J. M. (1955). The movements and migrations of mullet (Mugil cephalus L.). Australian 

Journal of Marine and Freshwater Research 2, 328-347. 

Thorrold, S. R., Jones, C. M., Campana, S. E., McLaren, J. W., & Lam, J. W. H. (1998). Trace 

element signatures in otoliths record natal river of juvenile American shad (Alosa 

sapidissima). Limnology and Oceanography 43 (8), 1826-1835. 

Tomás, J., Augagneur, S., & Rochard, E. (2005). Discrimination of the natal origin of juvenile allis 

shad (Alosa alosa) in the Garonne - Dordogne basin (south-west France) using otolith 

chemistry. Ecology of Freshwater Fish 14, 185-190. 

Tzeng, W. N., Shiao, J. C., & Iizuka, Y. (2003). Use of otolith Sr:Ca ratios to study the riverine 

migratory behaviors of Japanese eel Anguilla japonica. Marine Ecology Progress Series 245, 

213-221. 

Volk, E. C., Blakley, A., Schroder, S. L., & Kuehner, S. (2000). Otolith chemistry reflects migratory 

characteristics of Pacific salmonids: Using otolith core chemistry to distinguish maternal 

associations with sea and freshwaters. Fisheries Research 46 (1-3), 251-266. 

Wang, C. H., Tzeng, W. N., & You, C. F. (2006). Geographic variation of otolith elemental 

composition in the juvenile grey mullet (Mugil cephalus) collected from Taiwanese estuaries. 

In press , 


6. Author addresses 

J. Tomás 

K. Anastasopoulou 

P. Berrebi 

P.D. Cowley 

A. Darnaude 

P. S. Diouf 

J.-D. Durand 

Postal: IRD - UR 070, UMR 5119, Laboratoire ECOLAG, Université Montpellier 2, 

cc 093, Place E. Bataillon, 34095 Montpellier Cedex 5, France 

Email javier.tomas@mac.com 

Postal: Hellenic Centre for Marine Research, Institute of Marine Biological 

Resources, Aghios Kosmas, 167 77 Hellinikon, Athens, Greece 

Email kanast@ath.hcmr.gr 

Postal: UMR 5554 Institut des Sciences de l'Evolution, Equipe Génétique et 

Environnement / Métapopulations, Conservation et Co-évolution Université 

Montpellier II, CC 065. Place E. Bataillon 34095 Montpellier Cedex 5, France 

Email berrebi@univ-montp2.fr 

Postal: South African Institute for Aquatic Biodiversity, Private Bag 1015, 

Grahamstown, 6140, South Africa 

Email p.cowley@ru.ac.za 



Email audrey.darnaude@univ-montp2.fr 

Postal: WWF WAMER, Sacré Coeur 3, n° 9442, Dakar, Senegal 

Email psdiouf@wwfsenegal.org 

Postal: IRD UR 070 RAP, Route des Hydrocarbures, BP 1386 Bel Air, Dakar, 

Senegal 

Email Jean-Dominique.Durand@ird.sn 

D. Flores-Hernandez Postal: Centro EPOMEX, Universidad Autónoma de Campeche, Av. Agustín Melgar 

y Juan de la Barrera, Apdo. Postal 520, 24030 Campeche, Mexico 

Email doflores@mail.uacam.mx 

B. Morales-Nin 

W.N. Tzeng 

C.-H. Wang 

A.K. Whitfield 

J. Panfili 

Postal: Instituto Mediterráneo Estudios Avanzados (CSIC/UIB), 

Miguel Marqués 21, 07190 Esporles, Islas Baleares, Spain 

Email ieabmn@uib.es 

Postal: Institute of Fisheries Science, National Taiwan University, No. 1, Sec. 4, 

Roosevelt Road, Taipei, 10617 Taiwan, Republic of China 

Email wnt@ntu.edu.tw 

Postal: Institute of Fisheries Science, National Taiwan University, No. 1, Sec. 4, 

Roosevelt Road, Taipei, 10617 Taiwan, Republic of China 

Email chwang@mail.ncku.edu.tw 

Postal: SAIAB, Private Bag 1015, Grahamstown 6140, South Africa 

Email A.Whitfield@ru.ac.za 



Email panfili@ird.fr 


7. Annexure 1 – WS3 Agenda 

WORKSHOP 3 


Main Uses of the Grey mullet as Indicator of 

Littoral environmental changes 

2006 - 2009 

Agenda 

26 - 28 September 2007 

INCO-CT-2006-026180 MUGIL Deliverable 4 – Feb. 2008

Homepage: 

www.mugil.univ-montp2.fr 

Coordinator: 

J. Panfili (IRD) 

Contents 

WORKSHOP 3 


Main Uses of the Grey mullet as Indicator 

of Littoral environmental changes 

2006 - 2009 

26 - 28 September 2007 

Objectives 

MUGIL will elaborate guidelines for migration 

studies (otolith microchemistry, fishery data, 

genetics, telemetry). All participants will be 

compelled to using the same protocols. 

European 

Commission 

INCO-CT-2006-026180 


Page 

1. OBJECTIVES OF MUGIL WS3 ON MIGRATION ······················································································ 2 

2. AGENDA······················································································································································· 2 

3. LIST OF PARTICIPANTS····························································································································· 5

1. Objectives of MUGIL WS3 on migration 

Dear MUGIL colleagues, 

The third MUGIL Workshop (WS3), related to Mugil migration studies, will be held at the 

ECOLAG laboratory (Montpellier 2 University) in Montpellier, France, 26-28 September 2007 

(host: Jacques Panfili). As you all know, this WS3 was initially scheduled to be hosted by 

IMEDEA in Palma de Mallorca, Spain. This change of host responds to unexpected logistic 

problems at IMEDEA. Nonetheless, we have ensured, in coordination with IMEDEA, that the 

change of host will not in any way compromise the success of this WS. 

WS3 will be focused on the topic of migration, and more precisely on approaches to follow 

individual movements of mugilids. The objective of studying migration within MUGIL is to 

assess the differences between ontogenetic migrations and migrations caused by other 

factors (environmental conditions, anthropogenic causes). It will start with a synthesis on 

current and past results obtained in research studies on different fields of interest: otolith 

microchemistry, telemetry, genetics and fishery data. WS3 will more precisely focus on the 

application of otolith microchemistry to study Mugil cephalus migrations. WS3 will thus be 

dedicated to agree on a standard protocol for otolith dissection, preparation and processing 

for bulk and surface analysis as well as on data treatment. The synthesis and personal 

experiences will give the basis to decide on one eligible method of processing and analysis. 

Finally, WS3 will provide a deliverable permitting to choose a standard protocol for otolith 

processing as well as methods, instruments and recommendations on the use of these tools. 

At the end MUGIL will elaborate guidelines for migration studies and at least all participants 

will be compelled to using the same protocols. 

This meeting will also be the occasion to start discussions on the preparation of a new 

proposal to be submitted to the EU: 

We are looking forward to seeing you in Montpellier. 

Yours sincerely, 

Javier Tomás & Jacques Panfili 

2. Agenda 

Wednesday 26 September 2007 – Overview of available methods 

9h - 10h00 

- M. Troussellier (director of ECOLAG laboratory, UM2). Welcome. 

- J. Panfili. Information on MUGIL WS3 accommodation. 

- Update of MUGIL internet web-pages. 

10h00 - 10h30 

- J.-D. Durand / P. Berrebi. Application of genetic markers to the study of Mugil 

cephalus migration. 

10h30 - 10h45 

Coffee break

10h45 - 12h30 

- P. Cowley. The use of telemetry in studies of Mugil cephalus migration. Insights into 

technology and methods. 

- J. Tomás. Otolith microchemistry: principles and application to the analysis of fish 

migration. 

- J. Tomás. Instruments available in otolith microchemistry. Which method for which 

approach? 

12h30 - 14h00 

LUNCH at Montpellier 2 University 

14h00 - 16h00 

- C.-H. Wang. Application of otolith microchemistry to the study of Mugil cephalus 

migration in Taiwanese waters. 

- A. Darnaude. Application of otolith microchemistry to the study of Mugil cephalus 

migration in Corsica waters (Mediterranean Sea). 

- W.-N. Tzeng. Mugil cephalus migration around Taiwanese waters assessed through 

fishery data. 

- A.K. Whitfield. Historical movements and migrations of Mugil cephalus in the St Lucia 

Lake system, South Africa. 

- K. Anastasopoulou. Movements of Mugil cephalus in the Hellenic waters. 

- Discussion on methods for migration studies on Mugil cephalus. 

16h00 - 16h15 

Coffee break 

16h15 - 17h00 

- Otolith dissection of Mugil cephalus from the Mediterranean specimens for 

microchemistry analysis. 

DINNER free 

Thursday 27 September 2007 – Otolith microchemistry 

9h - 10h30 

- Otolith preparation of samples of Mugil cephalus collected within MUGIL for bulk and 

surface analysis (embedding, sectioning). Overview of standard protocols. 

10h30 - 10h45 

Coffee break 

10h45 - 12h30 

- Bulk analysis of samples of Mugil cephalus collected within MUGIL using SB-ICPMS 

(Solution Based Inductively Coupled Plasma Mass Spectrometry) at the ISTEEM 

laboratory (UM2). 

12h30 - 14h00 


14h00 - 16h00 

- Bulk analysis of samples of Mugil cephalus collected within MUGIL using SB-ICPMS 

(Solution Based Inductively Coupled Plasma Mass Spectrometry) at the ISTEEM 

laboratory (UM2). 

- Data treatment: from raw data to clean data (limits of detection, precision of 

measurement and standards). 

16h00 - 16h15 

Coffee break 

16h15 - 17h00

- Discussion on otolith microchemistry protocols for migration studies on Mugil 

cephalus. 

DINNER free 

Friday 28 September 2007 

9h - 10h00 

- Synthesis on the application of otolith microchemistry to migration studies of Mugil 

cephalus. 

10h00 - 10h30 

- Comparison of the different methods to study migration within MUGIL: otolith 

microchemistry, telemetry, genetics, fishery data, biomarkers. How can these 

methods be made complementary within MUGIL? 

10h30 - 10h45 

Coffee break 

10h45 - 12h30 

- WS3 report. 

12h30 - 14h00 


14h00 - 16h00 

- Working group on the preparation of a new proposal to be submitted to the EU. 

16h00 - 16h15 

Coffee break 

16h15 - 17h00 

- WS3 report finalisation 

MUGIL DINNER (with confirmation) 

Saturday 29 September 2007 

9h00 - 17h00 

- Visit of the Sète fishing habour and the lagoons of the area.

3. List of participants 

Name Institute Country MUGIL 

N° 

Date of 

Arrival 

Date of 

Departure 

Jacques PANFILI IRD France 1 25/09/07 29/09/07 

Javier TOMÁS IRD Spain 1 25/09/07 29/09/07 

Jean-Dominique DURAND IRD France 1 25/09/07 29/09/07 

Laurent VIGLIOLA IRD France 1 25/09/07 29/09/07 

Catherine ALIAUME CNRS-UM2 France 4 25/09/07 29/09/07 

Patrick BERREBI CNRS France 4 25/09/07 29/09/07 

Audrey DARNAUDE CNRS France 4 25/09/07 29/09/07 

Pape Samba DIOUF WWF-Wamer Senegal 5 23/09/07 30/09/07 

Beatriz MORALES NIN IMEDEA UIB Spain 10 25/09/07 29/09/07 

Wann-Nian TZENG IFS-NTU Taiwan 12 24/09/07 29/09/07 

Chia-Hui WANG IFS-NTU Taiwan 12 24/09/07 29/09/07 

Katerina ANASTASOPOULOU HCMR Greece 13 25/09/07 29/09/07 

Alan WHITFIELD SAIAB S. Africa 14 25/09/07 30/09/07 

Paul COWLEY SAIAB S. Africa 14 25/09/07 30/09/07 

Meeting venue 

Laboratoire ECOLAG - UMR 5119 - Université Montpellier 2 - cc 093 - Place E. Bataillon - 

34095 Montpellier Cedex 5 - France 

Tel +33 (0)4 67 14 41 23 (J. Panfili) / +33 (0)4 67 14 37 05 (secretary) 

Web: http://www.ecolag.univ-montp2.fr/ 

Hotel venue 

HOTEL IBIS MONTPELLIER COMÉDIE - Le Triangle - allée Jules Milhau - 

34000 Montpellier - France 

Tel +33 (0) 4 99 13 29 99 

Fax +33 (0) 4 67 58 77 50 

Web: http://www.ibishotel.com/ibis/fichehotel/gb/ibi/0592/fiche_hotel.shtml 

Transport = 5-10 min walk from TGV train station - 15-20 min drive from the airport (taxi fee = 15-25 €)

Mugil migration studies - MUGIL PROJECT - Université Montpellier 2

You also want an ePaper? Increase the reach of your titles

Delete template?

Save as template?