Cephalopods Age Determination by Statoli - FAO

7/28/2019 Cephalopods Age Determination by Statoli - FAO

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ADRIAMEDSCIENTIFIC COOPERATION TO SUPPORT

RESPONSIBLE FISHERIES IN THE

ADRIATIC SEAGCP/RER/010/ITA

GCP/RER/021/EC

Cephalopods Age Determination by Statolith Reading:

a Technical Manual

AdriaMed Technical Documents No. 22 Rome (Italy), November 2007

GCP/RER/010/ITA/TD-22


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The conclusions and recommendations given in this and

in other documents in the Scientific Cooperation to

Support Responsible Fisheries in the Adriatic Sea Project

series are those considered appropriate at the time of

preparation. They may be modified in the light of further

knowledge gained in subsequent stages of the Project. The

designations employed and the presentation of material in

this publication do not imply the expression of any

opinion on the part of FAO or MiPAAF or EC concerning

the legal status of any country, territory, city or area, or

concerning the determination of its frontiers or

boundaries.


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Preface

The Regional Project Scientific Cooperation to Support Responsible Fisheries in theAdriatic Sea (AdriaMed) is executed by the Food and Agriculture Organization of the

United Nations (FAO) and funded by the Italian Ministry of Agriculture Food and Forestry

Policies (MiPAAF) and since 2007 from the Directorate General for Fisheries and Maritime

Affairs of the European Commission.

AdriaMed was conceived to contribute to the promotion of cooperative fishery management

between the participating countries (Republics of Albania, Croatia, Italy, Montenegro and

Slovenia), in line with the Code of Conduct for Responsible Fisheries adopted by the UN-

FAO.

Particular attention is given to encouraging and sustaining a smooth process of international

collaboration between the Adriatic Sea coastal countries in fishery management, planning and

implementation. Consideration is also given to strengthening technical coordination between

the national fishery research institutes and administrations, the fishery organizations and the

other relevant stakeholders of the Adriatic countries.

FAO-AdriaMed Project HQ

FAO FIFM

Viale delle Terme di Caracalla00153 Rome, Italy

Tel: ++39 06 570 55467

Fax: ++39 06 570 55188

e-mail: [email protected]

URL: http://www.faoadriamed.org


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GCP/RER/010/ITA Publications

The AdriaMed Project publications are issued as a series of Technical Documents

(GCP/RER/010/ITA/TD-00) and Occasional Papers (GCP/RER/010/ITA/OP-00) related to

meetings and research organized by or conducted within the framework of the Project.

Occasionally, relevant documents may be translated into national languages as AdriaMed

Translations (GCP/RER/010/ITA/AT-00).

Comments on this document would be welcomed and should be sent to the Project

headquarters:

AdriaMed Project

FAO FIFM

Viale delle Terme di Caracalla

00153 Roma

Italy

[email protected]

For bibliographic purposes this document

should be cited as follows:

Ceriola, L. and Milone, N. 2007. Cephalopods Age Determination by Statolith Reading: aTechnical Manual. Scientific Cooperation to Support Responsible Fisheries in the Adriatic

Sea. GCP/RER/010/ITA/TD-22.AdriaMed Technical Documents, 22: 78pp.


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Preparation of this document

This document is the final version of the material used for the preparation of two trainingcourses on age determination of cephalopods by statolith readings organized in 2006 and in

2007 by the FAO Projects AdriaMed and MedSudMed. These notes aim at providing an

introductory treatment of some of the basic topics of age determination using statoliths. The

notes serve as a reference guide to the methods used during the training courses. These notes

are primarily for the scientists involved in the AdriaMed and MedSudMed Project research

activities related to the appraisal of fisheries resources, they can also be of interest for

students and professionals of fisheries research. However, a comprehensive introduction to

the methodologies to be applied is outside the scope of this publication. This manual should

be considered a further integration to the information reported by Jereb et al. (1991) in view

of the research carried out and the progress made thereafter. It provides an updated and easily

accessible tool to facilitate the expansion of statolith analysis in the Mediterranean where, in

spite of the great importance of cephalopods in fisheries, research on this mollusc class still

has to be developed entirely. Interested readers may find the literature given in Chapter 9,

useful for in-depth treatment of the topic and the Glossary annexed to the document.

Acknowledgements

Special thanks are due to Ms Patrizia Jereb for her valuable scientific advice which has led to

the completion and the improvement of this document. Many thanks are also due to

Dr George Jackson for his support and the technical information provided. The assistance of

Ms Caroline Bennett in the preparation of this document is gratefully acknowledged.

Ceriola, L. and Milone, N.

Cephalopods Age Determination by Statoliths Reading: a Technical Manual.

AdriaMed Technical Documents. No.22. GCP/RER/010/ITA/TD-22, Rome, 2007: 78 pp.

ABSTRACT

The main objective of this document is to provide a useful guideline to the age

determination methodologies applied to cephalopods using statolith reading. The document

is the result of the training activities on this issue organized by the FAO regional ProjectsAdriaMed and MedSudMed in 2006 and 2007. A selection of methods to extract, prepare

and examine statoliths is described in detail. A brief introduction describing the importance

of growth studies in cephalopods, as well as the function, internal structure and shape of

statoliths is also included. Chapter 1 contains the background information. Chapter 2

describes the terminology used and illustrates the position, morphology and function of

statoliths in cephalopods. In Chapter 3 the equipment, materials and the procedure used to

extract, clean and prepare statoliths for reading, and the methods for growth increment

counting (statoliths reading) are described. Finally in Chapter 4 the potential results of the

age determination by using statolith analysis are illustrated. A Glossary is also included in

the manual.


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Table of Contents

Acknowledgements............................................................................................................. iv

Table of Contents ................................................................................................................ v

1. Background information................................................................................................. 1

1.1 AdriaMed and MedSudMed training activities........................................................ 1

1.2 Cephalopods in the marine ecosystem ...................................................................... 1

1.3 Cephalopod growth estimates ................................................................................... 4

1.4 Aim of this document ................................................................................................ 8

2. Cephalopod statoliths...................................................................................................... 9

2.1 Introduction............................................................................................................... 9

2.2 Shape and current terminology ................................................................................ 9

2.3 Structure and microstructure ................................................................................. 123. Statolith analysis ........................................................................................................... 21

3.1 Equipment and materials........................................................................................ 21

3.1.2.Collecting data sheet......................................................................................... 22

3.1.3 Measuring and dissecting kit............................................................................ 23

3.1.4 Cleaning and storing......................................................................................... 24

3.1.5 Mounting and grinding..................................................................................... 24

3.1.6 Counting (statolith reading) ............................................................................. 27

3.2 Procedure and techniques....................................................................................... 28

3.2.1 Extraction.......................................................................................................... 28

3.2.1.1 Chemical method ....................................................................................... 29

3.2.1.2 Surgical method ......................................................................................... 293.2.2 Cleaning and storage ........................................................................................ 32

3.2.2.1 Cleaning...................................................................................................... 33

3.2.2.2 Storage........................................................................................................ 33

3.2.3 Mounting and grinding..................................................................................... 35

3.2.3.1 Mounting .................................................................................................... 36

3.2.3.2 Grinding ..................................................................................................... 38

3.2.4 Counting............................................................................................................ 42

3.2.4.1 Direct counting........................................................................................... 45

3.2.4.2 Image analysis system ................................................................................ 47

4. Some possible outcomes ................................................................................................ 49

5. References...................................................................................................................... 50Glossary............................................................................................................................. 61

Annex A: Data sheet.......................................................................................................... 65

Annex B: Statoliths extraction from fresh animals (Sepioidea, Teuthoidea and

Octopoda) .......................................................................................................................... 66

Annex C: Ground statoliths.............................................................................................. 74


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1.Backgroundinformation

1.1 AdriaMed and MedSudMed training activities

The training component has been central to the AdriaMed and MedSudMed Projects since

their outset. During the AdriaMed and MedSudMed Coordination Committeemeetings, the

necessity to support and develop the Project component on training as a key part of the work

programme was emphasised. Other regional experts meetings organised by the Projects over

the years, also highlighted this requirement. Several training activities were therefore planned

and organized in agreement with the needs expressed by the research institutions involved in

the Projects activities. For AdriaMed this was mainly achieved through the Project Working

Group on Demersal Fisheries Resources; in particular several training courses were

organized by AdriaMed to enhance the standardization of the methodologies used for the

appraisal of demersal resources at Adriatic level. An annual, international bottom trawlsurvey was organized among the eastern countries of the Adriatic Sea, supported by the

AdriaMed Project, for the joint appraisal of demersal resources in the region.

However, it became increasingly clear from the Adriatic experts that national expertise

needed to be developed and improved within the region, and particularly in cephalopod

appraisal methods. The same request was expressed by the research institutions participating

in the FAO MedSudMed Project (that focuses on the Central Mediterranean).

Consequently, a first training course was organized in the framework of MedSudMed

activities in July 2006, a second training course was then organized by AdriaMed in May

2007 in Split (Croatia) in order to provide basic knowledge on age reading techniques, inparticular concerning the reading of cephalopod statoliths. The didactic material prepared and

used during these training exercises has therefore been drawn on to prepare this manual.

These notes aim to provide an introductory treatment of some of the basic topics of age

determination using statoliths. The notes serve as a reference guide to the methods used

during the training courses and are primarily for the junior scientists involved in the

AdriaMed and MedSudMed Project research activities related to the appraisal of the fisheries

resources. They can also be of interest for students and professionals of fisheries research.

1.2 Cephalopods in the marine ecosystem

Cephalopods play an important role in the marine ecosystems, both as predators and prey.

They are voracious, opportunistic and highly versatile predators of fish and invertebrates, and

also represent the most important prey category of several fish species and top predators such

as marine birds, whales and other marine mammals (for extensive reviews see Rodhouse and

Nigmatullin, 1996; Piatkowski et al., 2001; Boyle and Rodhouse, 2005).

In terms of biomass, on the basis of fisheries data, cephalopod production for the 1988-1991

period was estimated to be 1.88 percent of the total production necessary to sustain total

global fishery (i.e. 2.476 x 106t, mean annual net weight) (Pauly and Christensen, 1995, in

Boyle and Rodhouse, 2005). Human fishery for cephalopods has continued to rise since then


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and some information on their biomass has been added due to improved knowledge on the

consumption of cephalopods by natural predators.

In the Southern Ocean, based on an estimate of the consumption of cephalopods by

vertebrate predators (30 x 106t, Rodhouse et al., 1994a), a standing stock of squid biomass ashigh as 100 million tonnes was calculated. Further estimates of the whole cephalopod

standing stock biomass reached values between 193 and 375 million tonnes (Rodhouse and

Nigmatullin, 1996), and the average annual cephalopod consumption by sperm whale alone

was estimated to reach 267 x 106t (Clarke, 1996). According to these estimates, the

application of Pauly and Christensens approach (1995) to more recent data would lead to

considerably higher values. Although cephalopod incidence in predators diet may be over-

estimated (e.g. Santos et al., 2001) and fishery data may be not entirely accurate, if

considering that top predators harvest squids and fishes generally not available to human

fishery (e.g. Trites et al., 1997), values of total cephalopod biomass in the worlds oceans up

to 500 x 106t (Voss, 1973) may not be so unlikely. Being subdominant predators that tend to

increase in biomass when other species become depleted, and having been estimated to

consume between 2 and 4 x 109t of food annually (Rodhouse and Nigmatullin, 1996), it is

now clear that cephalopods are a dominant component within the marine ecosystem and that

ultimately fluctuation in their abundance may seriously affect the abundance of their

predators and prey populations (e.g. Jereb et al., 2005).

Originally not included among the economically important fishery resources, cephalopods

have gained increasing attention in the past decades as an alternative to more traditional

marine resources, and their importance as a source of protein for human consumption is

likely to continue to increase in the future (Caddy and Rodhouse, 1998; Piatkowsky et al.,

2001; Rosa et al., 2002; 2005). Due to a steady increase in cephalopod capture production

during the last 30 years, from about 1 to around 3.5 million Mt (FAO, 2006) (Figure 1), the

fishery they support became one of the top invertebrate fisheries in the world.

Figure 1. World capture production for cephalopod (1970-2005).

In the Mediterranean Sea, total cephalopod capture production increased steadily until the

end of the 1980s, when a peak was reached (83 000 tonnes in 1988), followed by a decrease

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during the 1990s,after which the total commercial landings settled down to about 50 000

tonnes per year (FAO, 2006) (Figure 2).

Figure 2. Mediterranean capture production for cephalopods (1970-2004).

A similar scenario characterizes the Adriatic Sea, where a decreasing trend in total

cephalopod landings was recorded starting from the mid 1990s (FAO, 2006) (Figure 3).

Figure 3. Total capture production for cephalopod in the Adriatic Sea (1970-2004).

This decreasing trend, anomalous if related to the continuously increasing trends observed in

the other areas of the distributional range, was described in detail and tentatively commented

on (Jereb, 2002; Jereb and Agnesi, in press), but no sound explanation has been given to date.

Major characteristics of the cephalopod populations studied so far are the relatively short life

span, high growth rates and relatively early maturity, along with a remarkable physiological

capacity to respond to environmental changes by quickly adjusting these biological features

(e.g. Boyle, 1983, 1987; Boyle and Boletzky, 1966; Boletzky, 1994; Roberts et al., 1998;

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Dawe et al., 2001; Rodhouse, 2001; Rodhouse et al., 2001; ODor et al., 2002; Waluda et al.

2004). Their high plasticity in growth-rate, in particular, provides cephalopods with an

advantage in the competition with long-living late-maturing finfish species, as highlighted by

capture production trends in highly harvested regions (Caddy and Rodhouse, 1998;Balgueras et al., 2000; Pauly et al., 2001; Payne et al., 2006; Ungaro et al., 2006; Ceriola et

al., 2007). These unique physiological strategies make cephalopods key-species in regions

characterized by considerable environmental fluctuations or high fishing pressure (Trotsenko

and Pinchukov, 1994; Laptikowsky and Remelso, 2001; Zeidberg and Hamner, 2002;

Jackson and Domeier, 2003; Ceriola et al., 2006; Chen et al., 2006; Dawe et al., 2006;

Jackson et al., 2007) and may ultimately result in dramatic abundance fluctuations, due to

change in oceanographic conditions (e.g. ODor and Dawe, 1998; Dawe et al., 2000; Bendik,

2001; Dawe et al., 2001;Jackson and Domeier, 2003).

Worldwide concern on the level of exploitation of all marine resources was the driving force

to increase knowledge for assessment and management purposes. Attention to cephalopods

came late, but have proceeded continuously since then and fishery-related studies are rapidly

contributing to knowledge on the biology of many cephalopod species (e.g. Rodhouse et al.,

1994b; Payne et al., 1998; Rodhouse et al., 2001; Boyle and Rodhouse, 2005; Jereb and

Roper, 2005; McIntyre, 2006). Due to the importance of growth (and growth rates) within

cephalopods life cycle, the understanding of this key-process is one of the main targets of

many recent comprehensive studies focusing on cephalopods (e.g. Jackson and ODor, 2001;

Ragonese et al., 2002; Arkhipkin, 2004; Jackson, 2004).

1.3 Cephalopod growth estimates

Cephalopod growth can be estimated by applying both indirect and direct methods.

Indirect method

The indirect method consists in the analysis of the length frequency distributions (LFD)

observed in samples obtained from commercial landings or experimental surveys (e.g.

Mangold-Wirz, 1963; Sanchez, 1984;Jereb and Ragonese, 1995), as is the practice for fish.

This involves the identification and interpretation of the different modes present in the

usually polymodal LFD of the selected species. Often this analysis is subjective and the final

outputs can vary considerably according to the readers interpretation.

In cephalopods, most of the studies have been carried out on squids and several modes

usually resulted in the studied LFD. These have been alternatively interpreted either as the

result of the overlapping of different sub-cohorts (because of a protracted spawning season)

or as sub-year or year classes, with different resulting life-cycle duration estimates (for

reviews see Arkhipkin, 2004; Jackson, 2004).

The debate is still open, hoewever to summarize, LFD analysis is considered suitable for

discrete and well-identified spawning periods (generally corresponding to a unimodal LFD

per time interval) as in the case of many fish species, while it seems less appropriate when

applied to species that have a protracted spawning activity, such as many cephalopods.


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Direct methods

a) Rearing

A direct method to investigate growth in cephalopods consists in the observation of theanimals in aquarium, where the variation in size (both length and weight) over periodic time

intervals can be measured (Figure 4). This provides sound information on individual growth

rate at different ontogenetic stages and allows an understanding of the influence of abiotic

and biotic factors on the growth process (e.g. temperature and food ration, see for example

Jackson and Moltschaniwskyj, 2001a; Villanueva 2000a,b; Forsythe et al., 2001; Villanueva

et al., 2003; Forsythe, 2004; Chung and Lu, 2005). However, several problems still limit a

wide application of this method. To date, not many cephalopod species have been

successfully reared in captivity for the entire life cycle, mostly due to a considerably high

mortality rate at the early stages. Thus, information on growth of reared animals may not

cover the whole life span, referring instead to a fraction of it (Arkhipkin, 2004). In addition,

uncertainty about the correspondence of growth in captivity to that in the wild remains, as it

is not possible to perfectly reproduce natural environmental conditions in an aquarium. In

spite of these negative constraints, studies performed on reared animals helped to understand

growth performance in several species and were used to validate results obtained by applying

other ageing methods (e.g. to validate the time interval necessary for the formation of growth

increments in statoliths and gladii) (Jackson, 1994; Arkhipkin, 2004; Jackson, 2004)

Figure 4. Cephalopod rearing in captivity: a-b eggs maintenance, c adults rearing. From CephBase

(http://www.cephbase.utmb.edu/)

b) Tagging and recapture experiments in the wild

Tagging cephalopods in the wild and recapturing them, by knowing the time interval (i.e. the

time that passes from the tagging and the recapture), offers another direct way to measuregrowth of this group of species. This method has been used for a long time, from the very

early experiments in the late 1920s (for review see Nagasawa et al., 1993), until the recent

multi-species electronic Tagging of Pacific Pelagics (TOPP), a pilot program of the Census

of Marine Life (Block et al, 2002). Mainly applied to investigate vertical and horizontal

migrations and geographic distribution of commercially important ommastrephid

populations, this methodology also allows for the collection of important information on the

biology, physiology, ecology and stock identity of the investigated populations (Nagasawa et

al., 1993). Several tags can be used, depending mainly on the individual size range of the

species investigated, on the research targets and on the amount of funding available. Tags

more commonly applied to cephalopods are shown in Figure 5. Also in this case, however,

ab

c


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unavoidable disadvantages have limited the use of the methodology to investigate growth in

cephalopods and in squids in particular. Indeed, many squids are highly migratory animals

and their recapture rates are very variable (for reviews see Nagasawa et al., 1993; Murata and

Nakamura, 1998; Sauer et al., 2000; Markaida et al., 2005; Starket al., 2005; Semmens etal., 2007); in addition, the tagging process itself may affect animals mantle integrity and

swimming capability and eventually their survival, such that, in general, juveniles and small

size specimens cannot be tagged (e.g., Semmens et al., 2007).

Figure 5. Tags generally used to mark small fish and cephalopods: (a) Spaghetti tag; (b) Dart tag; (c) T-bar

anchor tag.

c)Hard structures analysis

All the hard structures in cephalopods soft bodies, with the exception of the chitinous

suckers rings, have the potential to memorise ontogenetic events through the formation of

periodical marks, or growth increments (see Arkhipkin, 2005, for a review) (Figures 6,7).

These structures include statoliths (Young, 1960; Clarke, 1966), gladii (La Roe, 1971),

mandibles (Clarke, 1965) crystalline lens of the eye (Williams, 1909) and cuttlebones (Choe,

1963).

Figure 6. Schematic illustrations of a cephalopod, showing the location of hard structures potentially suitable

for age and growth studies: Statoliths; Bones: Gladius and Sepion; Beak; Crystalline lens.

(a) (b) (c)

Beak

Statolith

Cuttlebone

Gladius

Crystalline lens


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Figure 7. Sucker rings (SR), probably the only hard structure in cephalopods bodies not bearing growth

increments.

Statoliths, in particular, are considered as true archives of cephalopod life cycles (Rhoads and

Lutz, 1980 in: Lipinski, 2001) and can be used to investigate several aspects of their

physiology, ecology and life style (e.g. Clarke and Maddock, 1988; Bizikov, 1991; Lipinski,

2001; Arkhipkin, 2005; Villanueva et al., 2007). Most important to the purpose of this

manual, all these structures have the necessary characteristics to be used as ageing tools: (i)

the presence of interpretable increment microstructures; (ii) the possibility to correlate these

microstructures progression/evolution with a regular and determinable time scale; (iii)

continuous growth at a measurable rate throughout the whole life span, especially in squids

(Beamish and McFarlane, 1983 in: Arkhipkin, 2005).

The first to notice periodical marks on statoliths was Young (1960) who studied the

statocysts in Octopus vulgaris. However, only in the middle of the 1960s was the role of

statoliths as recording structures recognized, when growth increments in the statolith

microstructure of several squid species were described (Clarke, 1966). Since then, several

studies have been carried out and outstanding progresses have been made in understanding

statoliths structure, function and their possible use to investigate cephalopod growth (e.g.Rodhouse and Hatfield, 1990; Jereb et al., 1991; Arkhipkin and Perez, 1998; Arkhipkin and

Bizikov, 2000; Jackson and ODor, 2001; Lipinski, 2001; Bettencourt and Guerra, 2000;

Arkhipkin, 2004; Jackson, 2004; Arkhipkin, 2005; Ceriola, 2007; Jackson et al., 2007,

Zumholtz et al., 2007a,b).

Other than for age and growth estimates, these small calcareous structures proved to be

useful for species identification (e.g. Clarke, 1978; Lipinski et al., 1993; Clarke, 1998;

Lombarte et al., 2006) and to investigate several aspects of cephalopod life history, including

age, hatching date and hatching period, growth rate, ontogenetic shifts, accidental events,

number of mating events, variations in environmental conditions, population structure,

SR


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systematic, population dynamic and life style (e.g. Jackson, 1994; Jackson and

Moltschaniwskyj, 2001b; Arkhipkin, 2003; Jackson and Domeier, 2003; Steer et al., 2003;

Villanueva et al., 2003; Forsythe, 2004; Chung and Lu, 2005; Semmens et al., 2007;

Villanueva et al., 2007; Zumholz et al., 2006, 2007c). Indeed, their potential as actual liferecorders was eventually acknowledged (Arkhipkin, 2005).

As for age determination, several problems still wait to be solved, calling for further research

and studies: the pillar hypothesis of the daily periodicity of growth increments (i.e. 1 ring = 1

day), was only partially validated and only in one case did the validation occur in the natural

environment (Lipinski et al., 1998); many species have not been studied yet; statolith

analysis is laborious and very time consuming; the number of samples is often limited,

making age and growth estimates challenging; changes in statolith structure and growth

increment deposition rate do occur along with the animals growth and maturity, as changes in

the increment width do, and biases related to statolith preparation and differences in the

methods of interpreting and enumerating increments have been widely acknowledged (e.g.

Lipinski and Durholtz, 1994; Gonzalez et al., 2000; Bettencourt and Guerra, 2001). In spite

of all these problems, to date statoliths microstructure analysis is the most widely used

method to investigate age and growth in cephalopods species, particularly squids (for reviews

see Arkhipkin, 2004; Jackson, 2004). Therefore, broad scale and collaborative studies on

their use are highly welcomed, to contribute to evaluating precision and to increase

consistency among investigators.

1.4 Aim of this document

To promote statolith analysis and gather further information on cephalopods and especially

on squid growth, in areas where statoliths are not yet currently used for age and growth

studies (e.g. many regions in the Mediterranean basin), practical guidelines are necessary. A

first manual on age determination by statolith analysis was published by Jereb et al. (1991)

and collects the proceedings of an international workshop held at the Istituto di Tecnologia

della Pesca e del Pescato of the Consiglio Nazionale delle Ricerche in Mazara del Vallo

(Sicily, Italy) in 1989, it includes a laboratory guide in which basic methods and techniques

used for statolith analysis are described.

This manual does not intend to replace the precious contribution provided by Jereb et al.

(1991), but rather to integrate the information therein reported, in view of the research carried

out and the progress made in the almost 20 years that have passed. The aim of this work is to

provide an updated and easily accessible tool to help spread the use of statolith analysis in the

Mediterranean areas and the Adriatic Sea in particular, where research and studies on

cephalopods are currently developing.


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2.Cephalopodstatoliths

2.1 Introduction

Cephalopod statoliths are paired calcareous structures associated with the sensory epithelia

(i.e. the macula statica princeps) located on the wall of the anterior chambers of the two

adjacent sac-like equilibrium organs called statocysts (Figure 8). First described by

Ovsjannikov and Kowalevsky (1867) and intensively studied thereafter (for brief reviews see

Budelmann, 1990; Arkhipkin and Bizikov, 2000), statocysts are cavities of irregular shape,

located in the posterior/ventral part of the cranial cartilage and consist of two chambers

(anterior and posterior), partially separated by finger-like projections and filled with liquid.

They can be considered analogous to the vertebrate semicircular system (Stephens and

Young, 1978; Williamson and Budelmann, 1985) and have two receptor systems (i.e. the

gravity and the angular acceleration receptor system) which give the animal proper

information about its position and movement in the water and enable it to compensate its eye

movements (Budelmann, 1977). The complex macula-statolith is responsible for the

detection of gravity and a possible role of statoliths in the detection of angular accelerations

was recently hypothesized (Arkhipkin and Bizikov, 1998). The potential of the statocists to

detect vibration stimuli was also investigated and the capability to detect low-frequencies

sounds was recorded and commented (e.g. Hanlon and Budelmann, 1987, Hanlon and

Messenger, 1996).

Figure 8. Statocyst structure (anterior part) and position of the statoliths (from Arkhipkin and Bizikov, 2000

modified).

2.2 Shape and current terminology

Statolith shape is species-specific and varies greatly both in the same species, during the

ontogenesis from a simple droplet in paralarvae (Figure 9) to the differentiated statoliths in

STATOLITHS

STATOCYSTS


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all adult specimens (Arkhipkin, 2003), and among the different cephalopod groups,

octopods, squids and cuttlefishes (Figure 10).

Figure 9. Illex illecebrosus statoliths: A) from a newly hatched paralarval (0 day old); B) from a 7 days old

specimen (from Sakai et al., 2004).

Figure 9. Statoliths of (A) Loligo vulgaris, (B) Sepia officinalis, (C) Eledone moscata at the transmitting light

microscope.

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Clarke and Maddock (1988) suggested that statolith shape depends more on the evolutionary

relationships of cephalopods rather than on the statoliths function. Arkhipkin and Bizikov

(2000) confirmed this hypothesis for squids and sepioids and distinguished two main statolithmorphologies: the demersal, typical of near-bottom decapods, and the pelagic, typical of

all pelagic squids.

Teuthoid and sepioid adult statoliths are considerably different from octopod statoliths and

their external morphology is characterised by four main regions: the dorsal dome, the lateral

dome, the rostrum and the wing (Clarke, 1978) (Figure11). The wing, which is the area of

attachment to the statocyst wall (Dilly, 1976), has a dorsal and a ventral indentation,

separated by a structure called the spur. Statoliths are predominantly hard and translucent

except for the wing, which is softer and opaque white.

For convenience and clarity in the present document, to describe statolith orientation

(anterior, posterior, dorsal, ventral, medial, lateral, marginal) and sections (sagittal,

transversal, frontal), the nomenclature reported by Lipinski et al. (1991) is adopted (Figure

12).

Figure 11. Adopted terminology to describe the external morphology of the statolith: (a) lateral view, (b)

anterior views; LD = lateral dome; DD = dorsal dome; R = rostrum; W = wing; S = spur; LS = lateral spur.

From Arkhipkin, 2005 modified.

(a) (b)

DD

DS

LD

S

W

R


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Figure 12. Adopted terminology to indicate statolith orientation and section planes. Left: D = dorsal; V =

ventral; L = lateral; M = medial; A = anterior; s = sagittal; t = transversal; f = frontal; m = marginal. (From

Lipinski et al., 1991 modified). Right: (1) = lateral view/section; (2) = frontal view/section; (3) transversal

view/section. The shape and proportions in the figures are only indicative.

2.3 Structure and microstructure

Statoliths are composed of calcium carbonate (CaCO3) crystallised as aragonite, which lie

within a matrix of organic material that has been ascertained to be mucoprotein (Radtke,

1983). The aragonite crystals are about the 95% in weight of the whole structure, with the

mucoprotein matrix accounting for the remaining 5%. However, an inverse relationship was

observed between this proportion and the individual size/age: the quantity of organic matter

declines with age, i.e. statoliths become more calcified as the animal grows up (Arkhipkin

and Perez, 1998; Bettencourt and Guerra, 2000).

Statolith formation results from a process called biomineralization (i.e. the deposition of a

mineral structure in a living creature) that was recently reviewed and described in detail forLoligo vulgaris and Sepia officinalis by Bettencourt and Guerra (2000). These authors

suggested that the different biochemical composition of the statocyst endolymph found in the

two species is responsible for different crystallization processes, which results in a different

microstructure and, ultimately, in a different definition of the growth increments. In addition,

they formulated the working hypothesis that dark rings (rich in organic matter) are deposited

during daylight, while light rings (rich in CaCO3) during darkness.

The crystals of carbonate and the protein matrix are deposited around a starting point named

focus (Natsukari et al., 1988) (Figure 13), which usually consists of one single concretion

(Arkhipkin and Perez, 1998). Around the focus statolith development proceeds periodically

(1) (2)

(3)


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with the deposition of new strata (or growth increments) outside the pre-existing external

surface. This process continues during the individuals entire life.

Figure 13. Detail of the statolith microstructure in ommastrephids (left) and loliginids (right); the arrows point

to F, the focus, the initial point of the statolith.

The width of each growth increment (0.5-10 m) varies depending on the deposition rates

and, indirectly, on the environmental variations or the ontogenetic shifts experienced by

the animal during the lifespan; for example, hatching and the transition between ontogenetic

phases have direct consequences on the deposition process inducing a marked anomaly in the

formation of the growth increments (Arkhipkin, 2005). The understanding and interpretation

of the growth increment periodicity, that is the period of time necessary for the developmentof a complete increment, is one of the main problems concerning the use of statoliths for age

and growth studies (for reviews see Arkhipkin, 1991; Lipinski et al., 1991; Jackson, 1994;

Arkhipkin and Perez, 1998; Arkhipkin, 2004; Jackson, 2004). Increasing evidence suggested

that increments are formed with a daily periodicity and that their total number represents the

individuals age in days (e.g. Lipinski, 1978; Hurley and Beck, 1979; Hurley et al., 1979;

Jackson and Choat, 1992; Jackson, 1994; Arkhipkin, 2004). The hypothesis one increment =

one day was then validated for several species by means of different experimental methods

(for reviews see e.g. Arkhipkin, 1991; Arkhipkin, 2004; Jackson, 2004). Even though this

assumption is now widely accepted, it is dutiful to point out that results of statolith analysis

should be taken with caution and the derived growth estimates should be considered

putative in species for which this validation has not been carried out yet (Lipinski andDurholtz, 1994; Lipinski et al., 1998).

In general, statolith microstructure reflects its periodical formation: when observed in a

proper section a number of concentric rings (for comments on the terminology see Lipinski

et al., 1991; and Lipinski, 1993) become visible with the centre in the focus (Figure 14).

These rings (hereunder also referred to as growth rings or growth increments)

correspond to the statoliths growth increments. They are composed of the paired dark and

light growth layers produced over a 24h period (Figure 15) (e.g., Arkhipkin, 1991; Lipinski

F

10 m

F

5 m


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et al., 1991; Jackson, 1994; Bettencourt and Guerra, 2000; Sakai et al., 2004; Arkhipkin,

2005; Jackson et al., 2005).

Figure 14. Statolith microstructure; narrow and large growth increments are highlighted.

Four main growth zones or regions, each characterised by rings of different width, are

generally observed within the statolith microstructure of adult squids: the nucleus and the

postnuclear, dark and peripheral zones (Figure 16). Each zone is formed during different

ontogenetic phases and the shift between adjacent zones is usually related to ontogenetic

shifts (Arkhipkin, 2005).

Figure 15. Dark and light layers alternate in the statolith growth increments.

5 m

50 m


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Figure 17. Detail of the statolith microstructure in (A) ommastrephids and (B) loliginids; NR, the natal ring, is

highlighted.

The natal ring corresponds to the first growth increment in ommastrephid squids (e.g.

Radtke, 1983; Balch et al., 1988; Arkhipkin and Perez, 1998; Villanueva et al., 2003; Steer et

al., 2003) and probably in most oegopsid squids, as growth increments in this group are only

deposited after hatching (Arkhipkin, 2004, 2005). On the contrary, in myopsid squids growth

increment formation begins during the embryogenesis and the newly hatched statoliths

already have several embryonic increments inside the natal ring (e.g., Morris, 1991;

Villanueva et al., 2003; Jackson and Moltschaniwsky, 2001a; Jackson, 2004) (Figure 18).

NR (B)

NR

(A)


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Figure 18. Natal ring and some internal rings in the statolith microstructure; NR = natal rings; . IRs = internal

rings.

25

NR

IRs

NRIRs

50

NR IRs

50


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The postnuclear zone (Figure 19) is generally formed within the immediate post-hatching or

paralarval phase. This zone is well defined in Ommastrephidae which are characterised by

having a special paralarval phase, named rhynchoteuthion (Chun, 1903), in which bothtentacles are fused into a proboscis.

Figure 19. Details of the statolith microstructure: the arrows indicate the inner and the outer boundaries of the

postnuclear zone (PNZ).

This constitutes a very delicate phase in the squids life cycle (Dawe and Brodziak, 1998),

since the animals seem unable to attack prey (Boletzky and Hanlon, 1983) and they probably

feed on suspension material (ODdor et al., 1985). The transformation to predatory juveniles

may be critical, by causing temporary starvation (e.g. Froeman, 1984) which results in a

PNZ

PNZ


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decreasing growth phase, recorded as prominent check in the statoliths microstructure (e.g.

Laptikhowsky et al., 1993; Arkhipkin, 2005). Other Oegopsid squids do not have a distinct

paralarval phase; thus, such a distinct boundary between the postnuclear and the following

zone is not evident in their statoliths (e.g. Arkhipkin and Perez, 1998; Arkhipkin, 2005).

Both the subsequent dark and peripheral zones are characterised by regularly spaced growth

increments (Figure 20). The influence of several factors on the formation of these zones was

studied, and the transition from the dark to the peripheral region was alternatively attributed

to changes in the food regime, as well as to habitat shifts (e.g. Jackson, 1993; Arkhipkin,

1997; Arkhipkin and Perez, 1998; Arkhipkin, 2005).

Figure 20. Statolith microstructure main four regions: (F) focus, (PNZ) postnuclear zone, (DZ) dark zone, (PZ)

peripheral zone.

Conspicuously marked rings or checks (Lipinski et al., 1991) are often noticeable within

the statoliths microstructure (Figure 21). These prominent rings are assumed to reflect the

occurrence of specific events in the cephalopods life, and were associated to various

stressful episodes such as behavioural changes, migrations, starvations, thermal shocks,

mating etc. (e.g., Spratt, 1978; Kristensen, 1980; Lipinski, 1981; Arkhipkin and Murzov,

1986; Arkhipkin, 1988, Arkhipkin and Perez, 1998, Arkhipkin, 2005).

FPNZ

DZ

PZ


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Figure 21. Checks in the statolith microstructure of adult females, in response of mating event.

check

check


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3.Statolithanalysis

3.1 Equipment and materials

Statolith analysis is a multi-phase procedure, which includes statolith extraction, cleaning,

mounting (on an appropriate support), grinding and, eventually, reading (i.e. growth

increments counting). The following commonly used laboratory tools and specific materials

and equipments are required to proceed (Figure 22):

cephalopods samples data collectingsheets dissecting and measuring kits cleaning and storing kits mounting and grinding equipment ring counting (statolith reading) equipment

A detailed description of the necessary equipment is hereby provided and the most important

characteristics of specific materials (e.g. glue paper for grinding) are described and

commented on.

Figure 22. Equipment and materials necessary to collect and prepare the statoliths: cephalopod samples (a)

cephalopod samples; (b) dissecting kit; (c) lapping films; (d) microscope slide, (e) mounting medium; (f)

transmitting light microscope.

(a)

(b)

(c)

(d)

(e)

(f)


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3.1.1.Samples

Fresh, frozen or alcohol-preserved cephalopods can be used for statolith extraction and

studies (Figure 23). Statoliths will be considerably easy to find in the cephalic cartilage offresh animals and also in defrosted specimens, whilst it can be difficult to extract them from

specimens preserved for a long time (more than two years) in ethyl alcohol (Lipinski, 1980;

Dawe and Natsukari, 1991; Jackson and Choat, 1992). Specimens preserved in formalin are

not suitable for statolith studies because formalin deeply damages calcareous structures.

Figure 23. Fresh samples: Sepia officinalis (left);Illex coindetii (right)

3.1.2.Collecting data sheet

The preparation of the data sheets for data recording should be carefully planned. A single

data sheet that permits the recording of a wide range of information (e.g. sampling source

and collecting date, name of the recorder, morpho-biometric characteristics of the specimenconsidered, as well as all the necessary information to identify the individual to which the

statoliths belong) should be prepared and used for each specimen (Figure 24, Annex A)

Figure 24. Example of a data collecting sheet for biological studies.


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3.1.3 Measuring and dissecting kit

A complete kit to measure and dissect cephalopods is necessary (Figure 25). This includes a

ruler (1 mm) and a scale (at least to 0.1 g, but to 0.01 g is recommended) for bodymeasurements (length and weight), scissors, a surgical blade or a scalpel, surgical tweezers

and a dissecting needle, to dissect cephalopods and extract the statoliths.

When handling very small individuals (i.e. embryos or paralarval individuals; mantle

length < 1 mm), a dissecting microscope (or a stereomicroscope) with 40X - 50X total

magnification and very fine tweezers and dissecting needlesare necessary. Larger specimens

require progressively less magnification (10 X 20 X for specimens of approximately 2-

10 mm DML), whereas individuals with mantle length over 100 mm generally do not require

a microscope for dissection (e.g. Dawe and Natsukari, 1991;Villanueva, 2000a,b; Villanueva

et al., 2003).

Figure 25. Kit for specimens dissection and measurements: (a) capsulae, blazers and scissor; (b) surgical blade

and pins; (c) ichtyometer; (d) ruler (0.1 cm).

(d)

(a)

(b)

(c)


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3.1.4 Cleaning and storing

Distilled water, absolute ethanol, or other chemical solutions such as sodium hypochlorite(1% - 5%) and NaOH (10%), can be used to clean the statoliths. Ultrasonic cleaners can also

be used (e.g. Dawe and Natsukari, 1991; Chung and Lu, 2005).

To store the statoliths any plastic box or capsule, as well as oil-paper envelopes are

potentially useful, as statoliths can be stored dry; however ethanol (70% or 96%) gelatine or

glycerol can also be used as storing media (Rodhouse and Hatefield, 1990; Dawe and

Natsukari, 1991; Jackson and Yeatman, 1996). The use of a plastic 96-well immunoassay

microplates as illustrated in Figure 26 is recommended. A label recording the statoliths basic

references is also necessary, i.e. a link to associate the statoliths to the specimens from which

they were extracted. Generally a paper label will do.

Figure 26. Plastic 96-well immunoassay microplates

3.1.5 Mounting and grinding

Before grinding, statoliths have to be mounted on an appropriate support, which is generally

a microscope slide. Accordingly, to mount a statolith on a microscope slide, a dissecting

microscope (20 X 40 X magnitude) a surgical needle and a proper mounting medium (i.e.

a glue or paste) are necessary (Figures 27, 28). The choice of the mounting medium is very

important, as it should guarantee strong adhesion to glass surfaces, good viscosity, fastdrying and transparency.

Several glues are used as mounting media, all of them presenting both advantages and

disadvantages: the Canada balsam, which needs a long time to dry, but has a good viscosity

(Lipinski, 1978); plastic substances as the Lakeside 70 cement (Arkhipkin, 1991); some

transparent resins like the Protexx syntetic mountant (Dawe et al., 1985), which also require

some time to complete hardening, or the Polarbel 812 resin (Morris and Aldrich, 1985); and

thermoplastic cements, like the Crystal Bond

(Jackson, 1990) and the Buehler

(Zumholz et

al., 2006). The DPX (dibutyl-phtalate-polystirene-xylene), a mounting medium used for

histological studies, has also been used for statolith analysis with double function, glue and


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clearing agent, especially for small species or sub-adult individuals statoliths, which

generally do not require grinding (e.g. Jackson, 1989; Dawe and Natsukari, 1991; Jackson

and Choat, 1992; Jackson, 1994).

Figure 27. (a) Fragment of Crystal Bond 509, the mounting medium suggested; (b) hot plate; (c) microscope

slide.

Currently, one of the most frequently used media is the Crystal Bond

, which proved to be

an excellent choice as it is completely translucent, it is not fluorescent under UV irradiation,

melts at low temperatures and hardens relatively rapidly after removal from heat (Jackson,

1990; Jackson, 1994; Jackson et al., 2005). Several types of Crystal Bond

are

commercialised, with different melting temperatures and properties; among these, the CrystalBond

509 (softening temperature 71 C, flow point 135 C, viscosity at flow point 6000

cps) can be considered an ideal medium for statolith mounting (e.g. Jackson, 1990;

Bettencourt and Guerra, 2001). Therefore, when using the Crystal Bond

or any type of

similar thermoplastic cement for mounting statoliths, a hot plate capable to achieve the

melting temperature (110-130 C for Crystal Bond

) will also be necessary.

(a) (b)

(c)


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Figure 28. Dissecting microscope (stereomicroscope 10-40 X total magnification) and microscope slide. The

arrow points to a statolith placed on the microscope slide.

To grind statoliths, the use of several different types of sandpapers, each with a proper grit, is

recommended. Among recent available commercial sandpapers, lapping films gave good

results (e.g. Zumholz et al., 2006) and in this manual the use of a complete set of lapping

films with 30, 12, 5, 3 or 0.05 grades is recommended to achieve the best results from

grinding1

(Figure 29). For the grinding process, the coarsest lapping film will be used first,

the others will follow in sequence following a decrease in coarseness.

A flat and clean surface is also required as a basis for the grinding process. A glass slide

large enough to contain the lapping films can be considered an optimal solution, but any

other material (e.g. metal or plastic) with the same characteristics will do as well (Figure 30).

1The 3M

lapping film currently is one of the most used sandpapers.


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Figure 29. Lapping films for grinding, with different grit grades (in parenthesis).

Figure 30. To grind the statolith, a glass slide or any other flat and clean surface is needed as a support for the

lapping films.

3.1.6 Counting (statolith reading)

For the analysis of statolith microstructure (e.g. to count growth increments, or to measure

hatch rings) a transmitting light microscope with a total magnification between 100 X

400 X (generally ocular 10 X with micrometer, objectives 10 X, 20 X, 40 X) is generally

required, although a magnitude of 600 X will be needed for observation of embryonic or

paralarvae statoliths. In some cases, the potential need for even higher magnification (i.e.

600-1000X) has been highlighted (Arkhipkin, 2005), while the use of polarized light can

improve ring identification and counting (e.g. Dawe and Natsukari, 1991; Jackson and

Moltschaniwskyj, 1999; Jackson et al., 2003) (Figure 31). The use of a system capable of

displaying statoliths microstructure on a monitor is highly recommended. Standard systems

(30 m)

(12 m)

(5 m)(00.5 m)

(30 m)(12 m)

(5 m)(00.5 m)


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generally consist of a camera mounted on a microscope and connected to a computer

monitor.

In some cases, on the basis of the experience carried out in studies on fish otoliths (see

Campana, 1992 and Jearld, 1995 for reviews), a computer-based image analysis system(Campana, 1987) was used for statolith investigation (Arkhipkin, 1996; Gonzalez et al.,

1996). An automatic, or semi-automatic ring counting, through an image analysis system, can

also be carried out. However, due to the variation of the growth increment width even in the

same statolith, the automatic ring counting has to be considered with care and it will not be

described in this manual.

Figure 31. Transmitting light microscope with 200X-600X total magnification minimum (top left) and an image

analysis system, with a camera (oval) mounted on a microscope (right) connected to a monitor (down left).

3.2 Procedure and techniques

3.2.1 Extraction

Two different methods can be used for statolith extraction: surgical and chemical. The

surgical methods are highly recommended, however the chemical method has also been used

(e.g. Hurley and Beck, 1979), thus a brief description of the latter method is given as well.


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3.2.1.1 Chemical method

The method involves the extraction of the posterior-ventral portion of the cartilaginous skulls

and its dissolving in a proteolyitic enzyme solution (e.g. pepsin and sodium hypochlorite,Hurley and Beck, 1979; trypsine and boric acid, Amaratunga and Budden, 1982) within an

appropriate vial. After the cartilage has dissolved, the two statoliths can be collected from the

bottom of the vial.

In order to extract the skull, set the individual with the ventral side up and anterior side

toward, and remove the funnel and the skin beneath it. The ventral part of the cephalic

cartilage will be exposed and can be removed by severing it from the head.

The use of chemicals may apparently simplify the extraction, but is not suitable when the

statolith sampling occurs on board vessels, as secure conditions to store and handle

potentially dangerous chemical components are required. Furthermore the use of chemicals is

not appropriate to process large samples. In addition, chemicals may eventually damage thestatoliths (Dawe and Natsukari, 1991).

3.2.1.2 Surgical method

Techniques for surgical statolith extraction have been described in many of the publications

dealing with cephalopod age and growth, the most useful being reviewed by Rodhouse and

Hatfield (1990), Arkhipkin, 1991 and Dawe and Natsukari (1991). In this manual two

surgical techniques for statolith extraction are described.

1) Place the animal ventral side up and anterior side towards you, and cut the fusionbetween funnel and body from the anterior part with a surgical blade or a scalpel. After

cutting the fusion, the statocysts will be visible through the skin in fresh and defrosted

squids, even in large specimens. In octopods and sepioids the skin and muscles beneath

the funnel generally prevent the view of the statocysts, thus this tissue has to be removed

by means of horizontal/longitudinal cuts (along the sagittal plane) performed holding the

animal with one hand and flexing the head dorsally. Once the cephalic cartilage is

exposed, carefully remove thin vertical/transversal cartilage slices to cut off the anterior

side of the statocysts. While doing this, care should be taken not to force the surgical

blade too deeply or too far forward, to avoid damaging other parts of the head. This

could cause internal white fluids to flow over, which could cover the statocysts and

prevent location of the statoliths. Stop the cutting when the two statoliths become visible

in the anterior side of the statocysts (two distinct cavities will appear and the two

statoliths will be visible as clear particles in the middle of them). Once the statoliths are

exposed, pick them up with the surgical tweezers or with the aid of the scalpel/blade tip.

Generally speaking, care must be taken to handle the specimens gently and avoid

crashing or moving their heads too roughly, since statoliths may detach from the anterior

wall of the statocysts and drop into the bottom of the cavity. Some phases of statolith

extraction following this technique are shown in sequence below (Figure 32a-e) and in

Annex B.


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Figure 32. Some steps of a surgical technique for the statoliths extraction in Sepioidea: a) Set the individual

ventral side up and anterior side towards you; b) Remove the fusion between the funnel and the body from the

anterior side; c) After removing the tissue covering the statocysts, make one or more transversal cut in order to

expose the statoliths; 4) Stop the cutting when the two statoliths become visible in the anterior side of the

statocysts (two distinct cavities will appear and the two statoliths will be visible as clear particles in the middle

of them); e) Pick the two statoliths up with thin tweezers or with the tip of the surgical blade.

a)

b)


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Figure 32. Continued

c)

d)


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Figure 32. Continued

2) Place the animal ventral side up and sever the head in correspondence with the anteriorend of the mantle. As in 1), the statocysts will generally be visible in fresh squids and

the statoliths will appear as opaque white structures within them. Remove the anterior

cartilage wall by gently cutting away thin cartilaginous slices, until the two maculae

with their statoliths are visible and the statoliths exposed. Collect them gently with the

surgical tweezers or with the aid of the scalpel/blade tip.

Both procedures are also suitable to process very small specimens as embryos or paralarvae,

and generally animals with dorsal mantle length < 10 mm, under a dissecting microscope; inthis a case very thin sharpened blade or scalpel and tweezers are needed.

3.2.2 Cleaning and storage

After extraction, statoliths should be prepared for analysis or stored. As the grinding process

removes surface material, cleaning the statoliths will not be necessary at all if they have to be

ground on both sides. On the contrary, thin and translucent statoliths (mainly from paralarvae

and embryos) need either to be ground on one side only (Dawe and Natsukari 1991;

Arkhipkin and Roa-Ureta, 2005; Villanueva 2000 a,b; Villanueva et al., 2003) or not to be

e)


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ground at all (e.g. Jackson and Yeatman, 1996; Jackson and Moltschaniwskyj, 2001b;

Jackson et al., 1997): in these cases cleaning is required. However, it is always preferable at

least to rinse the statoliths in distilled water and let them dry at room temperature before

storing them or proceeding with the analysis. As statoliths from very small specimens aredifficult to locate and handle, it is advisable to not remove them and store the entire specimen

until the actual analysis (e.g. Dawe and Natsukari, 1991).

3.2.2.1 Cleaning

Several methods are suitable to clean statoliths, including rinsing them either in distilled

water, ethanol (absolute or 75%), sodium hypochlorite, or NaOH solution (e.g. Rodhouse and

Hatfield, 1990; Dawe and Natsukari 1991; Jackson and Wadley, 1998; Steer et al., 2003), as

well as ultrasonic cleaning (Dawe and Natsukari, 1991; Natsukari, 1998; Chung and Lu,

2005) as summarized below.

a) Place the statoliths in a vial (or in the plastic 96-well immunoassay microplates previously

described) with distilled water or ethanol (75% - 100%) and let them dry at room temperature

(e.g. Jackson and Moltschaniwskyj, 1999; Arkhipkin, 1997; Sakai et al., 2004; Triantafillos,

2004; Jackson et al., 2007).

b) Set the statoliths on a glass surface (e.g. a microscope slide), rinse them in distilled water

and dehydrate them by rinsing in absolute ethanol (e.g. Jackson, 1989; Rodhouse et al.,

1994b; Jackson and Wadley, 1998; Bettencourt and Guerra, 2000).

c) Place the statoliths in a solution of sodium hypochlorite (from 1 % to 5 %) for differenttime-ranges, depending on the strength of the solution (the stronger the solution, the shorter

the time interval) and the size of the statoliths. Usually a few minutes will be necessary for

thick statoliths (e.g. Jackson, 1990; Jackson and Forsythe, 2002), less than 1 minute for small

and thin statoliths (e.g. Ikeda et al., 1999). After cleaning, rinse the statoliths in distilled

water and dehydrate them with 100% ethanol.

d) Place the statoliths in an ultrasonic cleaner for 1-3 minutes, then soak the statoliths in a

strong alkaline solution (e.g. 10 % NaOH) overnight (Natsukari, in Dawe and Natsukari

1991; Chung and Lu, 2005).

3.2.2.2 Storage

Statoliths are acellular mineralized structures, that will not decompose under relatively dry

conditions (Rodhouse and Hatfield, 1990). Accordingly, they can be stored dry in small glass

or plastic vials (e.g. Triantafillos, 2004; Jackson et al., 2007) (Figure 33). Storage in a

wet/liquid medium such as oil-paper envelopes (Arkhipkin, 2003) 70 % or 96 % ethanol is

also possible (e.g. Arkhipkin, 1997; Challier et al., 2002; Miyahara et al., 2006). However,

when storing statoliths in ethanol within closed capsules, these should be kept in cool

conditions. Indeed, high room temperature (e.g. > 30C) may cause an increase in the partial


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pressure of the ethanol determining the opening of the capsules and the loss of the statoliths.

The use of glycerol and gel capsules is also appropriate, but more expensive and less easy to

apply.

Several researchers store statoliths dry or in ethanol (70% to 96%) (Dawe and Natsukari,

1991) and in general no differences in growth increment visibility were found examining

statoliths kept dry and those preserved in alcohol for one year (Nastsukari, 1998). Therefore,

ultimately, the storage method will be selected based on personal preferences/choices.

However, to store statoliths dry in the plastic microplate already mentioned sheltering with

paraffin eventually closing with a proper cover is suggested (Figure 34).

As already stated, when having to store statoliths of embryos or paralarvae and/or animals

smaller than 30-50 mm ML, it is advisable to store the whole specimens instead of the

statoliths extracted. These animals should be preserved in 95 % ethanol, after a 15-20

immersion in 60 % ethanol (Arkhipkin, 1991). For long-term storage, ethanol should be

changed periodically (every 6 8 months).

Whatever system is used to store the statoliths, each pair has to be labelled separately, in

order to link the results of the analysis to the specimen to which the statolith belongs. It is

useful to underline that plastic microplates can be used both for cleaning and storing by

filling the holes with distilled water or 96% ethanol, setting the paired statoliths in the bottom

of one single hole and letting them dry at room temperature. Once the holes are completely

dry, cover them with paraffin and/or sticky-tape to prevent the statoliths from moving and

close the box with its cover.

Figure 33. Statoliths can be stored in plastic vials (dry or in ethyl alcohol) or in any other box.


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Figure 34. The use of a plastic 96-well immunoassay microplate to clean and store the statoliths (both dry or in

ethyl alcohol), is recommended.

3.2.3 Mounting and grinding

While in paralarvae or small juveniles of some species growth increments are visible without

grinding, large juveniles and adult statoliths require preparation before grinding, i.e. mounted

on an appropriate support (generally a microscope slide) and ground to be read.

Several procedures were described in Jereb et al. (1991) and used thereafter, either as they

were originally described, or slightly modified (e.g. Bettencourt et al., 1996; Arkhipkin,1997; Lipinski et al., 1998; Raya et al., 1999; Gonzlez et al., 2000; Bettencourt and Guerra,

2001; Challier et al., 2002; Villanueva et al., 2003).

In this volume a comparatively easy procedure is reported and to this extent the use of the

thermoplastic cement Crystal Bond (or equivalent) as a mounting medium is assumed, as

its characteristics gives several advantages during the statoliths preparation (e.g. it is

possible to melt a thermoplastic cement at any time during the procedure allowing for

changes in the statoliths position/orientation when necessary).


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3.2.3.1 Mounting

Mounting the statoliths on a microscope slide (or on any other substrate) in order to prepare

them for grinding is a relatively easy but extremely important process. It is mostly during thisphase that the plane and the area to be ground are decided, as well as the section of the

statolith that will be observed.

Accordingly, the statolith should be mounted with the proper orientation to enable the

operator to grind it following a well-defined plane; this should allow a section of a pre-

defined area to be obtained and create the best conditions for the observation of the

microstructure (Figure 35a,b) (e.g. Dawe and Natsukari, 1991; Dawe and Beck, 1997;

Linpinski et al., 1998; Hendrickson and Hart, 2006; Jackson et al., 2007).

Figure 35. Different statolith orientation on the microscope slide: a = concave side up; b = concave side down.

MM = mounting medium; F = focus

Several regions of the statolith can be used for ring observation, differing in increment width

as well as in increment sequence, which can be complete (i.e. without interruption or whiteareas from the natal ring to the statolith margin) or not.

Generally, a transversal section of the domes (dorsal and lateral domes) is used for ring

examination and counting (e.g. Jackson, 1989; Arkhipkin, 1991; Jackson and Choat, 1992;

Dawe and Beck, 1997; Bettencourt and Guerra, 2001; Quetglas and Morales-Nin, 2004;

Challier et al., 2005; Hendrickson and Hart, 2006; Ceriola, 2007; Jackson et al., 2007), even

though in some cases (i.e. for several loliginids and some oegopsids) a transversal section of

the rostrum is preferred (e.g. Natsukari and Komine, 1992; Bettencourt et al., 1996;

Arkhipkin, 1997; Raya et al., 1999; Arkhipkin, 2005).

F

MM

b

FMM

a


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In order obtain a transversal section of the statoliths, under a binocular dissecting microscope

(10-40X magnification), set a fragment2

of the thermoplastic cement in the middle of a

microscope slide and place the statolith close to this fragment with the anterior or the

posterior side up (the cement fragment size should be equivalent to that of the statolith;larger quantities also can be used, but this may increase the time of grinding and compromise

the final clearness of the ground surface). Place the microscope slide on the hot-plate

(previously warmed to around 120 C) to melt the thermoplastic resin and wait for a few

seconds. After the resin has melted, move the slide back under the microscope and set the

statolith in the middle of the melted cement (Figure 36).

Figure 36. Placing the statolith on the melted mounting medium

While placing the statolith in the melted cement, it is important to not turn it upside-down in

order to keep the predefined grounding plane. In addition, the statolith orientation should be

accurately checked under the binocular microscope, so as to have the possibility to modify it,

if necessary, while the cement is hardening.

If the statolith orientation is correct, let the cement harden; on the contrary, melt the

thermoplastic cement again by replacing the glass slide on the hot-plate and modify the

statoliths position or orientation.

The Crystal Bond hardening and the microscope slide cooling down will require 1 or 2

minutes at room temperature, after which the statolith will be ready for grinding.

The first orientation of the statolith varies depending on the species and on the side/s to be

ground (whether only one or both). When both the anterior and the posterior side of the

statolith have to be ground, beginning with the concave side down rather than with the

concave side up is generally a choice based on the personal experience of the author and in

some extent to the characteristics of the statolith/species. Likewise, when only one side of the

statolith needs to be ground the choice is related to the species to analyse. For example,

2 Small fragments of resin are obtained from larger pieces by using a strong needle.


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grinding the anterior side of the statolith results in a better ring identification in Illex

coindetii, whereas the opposite is true forLoligo vulgaris (Ceriola pers. observ.).

Help in taking this kind of decision may come from literature, as well as from personal

experience.

3.2.3.2 Grinding

Grinding is the most delicate phase of the whole procedure of statolith preparation, since the

rings identification depends on proper grinding and, ultimately, the results counting process

too.

Statoliths from embryos, paralarvae and small loliginids and sepioids do not need grinding,

as they are thin and translucent enough to allow a direct ring counting3

(e.g. Jackson, 1989;

Jackson et al., 1997; Jackson and Moltschaniwskyj 2001b).

The grinding procedure consists of two phases: the first of main grinding and the second ofpolishing. Generally a coarse waterproof sandpaper is used for the main grinding, i.e. to

remove the external side of the statolith, while a finer sandpapers are used to polish the

ground surface. It was noted that statolith reading improves when statoliths are ground with

sandpaper moistened with cold water (Arkhipkin, 1991), however also grinding with dry

sandpaper gives excellent results.

To grind the statolith, set four small squares/rectangles of lapping film (each of 12 x 8 cm

approximately) with 30, 12, 5 and 3 or 0.05 grades respectively on a glass surface (or any

other flat surface). Then, holding the microscope slide with the mounted statolith upside

down, start grinding the statolith on the first lapping film (30 ) with a very light pressure

and circular movements (Figure 37). The extent and intensity of grinding, as well as thegrinding plane should be monitored continuously by observing the ground surface using a

dissecting microscope (40 X magnification) and/or a transmitting light microscope (up to

100 X magnification).

Continue grinding until the focus becomes visible beneath the ground surface, when focusing

at the microscope or when the ground surface almost reaches the edge of the statolith.

Stop using the coarsest lapping film before the edge of the statolith is reached and continue

the process using the second lapping film (12 grades). Changing the sandpaper ensures

that: 1) grinding decelerates and does not proceed so far as to obliterate the focus or to

damage the external edge; 2) some of the scratches are removed from the ground area(Figures 38, 39).

3For such small statoliths, the use of clearing agents like Eukitt, Euparal xylene and DPX (dibutyl-phthalate-

polystyrene-xylene) for mounting (Jackson, 1989; Dawe and Natsukari, 1991) may improve the visibility of

growth increments. The same effect was obtained by Villanueva et al. (2003) by using tetracycline, a marking

agent, when incubatingLoligo vulgaris eggs.


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Figure 37. Sequence of grinding. In parenthesis the suggested lapping films grades

Figure 38. Subsequent phases in the grinding process; the clear area is the ground surface.

main grinding

polishing

(30 ) (12 ) (5 )

(30 )(12 ) (5 ) (0.05 )

(12 ) (5 ) (0.05 )

(5 )

(0.05 )


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Figure 39. Subsequent phases in the grinding process of a real statolith. (A) Only the dorsal and lateral domes

were ground; (B) Almost the whole surface was ground.

Continue the grinding process with the 12 grades lapping film until the ground surface,

slightly above the focus, reaches the most external point of the dorsal dome (the edge of the

statolith). After that, keep on grinding with the two finer lapping films (5 and 3 or 0.05

grades), in order to polish the ground area.

When the surface of the statolith is polished from all the scratches, examine the statolith at

the transmitting light microscope (10 X ocular and objectives 10 X, 20 X and 40 X or more)

in order to decide if the growth rings are adequately visible (i.e. properly identifiable) or if

grinding the opposite surface is necessary.

Ground area

(A)

Ground area

(B)


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If the statolith needs grinding on the second surface, it has to be turned upside down (that is

with the ground surface on the glass) and further ground on the second side. To turn the

statolith upside down, place the microscope slide on the hot plate (set at about 120 C) untilthe Crystal Bond melts, then place the microscope slide under the dissecting microscope

and change the statolith orientation with a needle.

By gently touching the statolith margin, this will remain attached to the tip of the needle and

it will be easy to raise the statolith and turn it upside down, with the flat surface on the glass.

Once the cement and the glass support have cooled down and the whole structure has

hardened (generally within 1 or 2 minutes), start with the grinding procedure again, exactly

as done for the first surface.

As a general rule, and especially for inexperienced researchers, when grinding the second

surface (but also for the first one) it is advisable to stop before reaching the statolith edge, to

avoid exceeding in the operation (overgrinding), as in over-ground statoliths is very difficult

to properly identify rings and to estimate the individuals age. A scheme of the grinding

process (set to obtain a transversal section) and the different levels of grinding are reported in

Figure 40 (convex side up) and Figure 41 (convex side down).

Figure 40. Longitudinal section of a mounted statolith (convex side up) to illustrate the grinding process as it

proceeds on both sides of the statolith. The dashed line indicates the grinding plane.


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Figure 41. Longitudinal section of a mounted statolith (convex side down) to illustrate the grinding process as it

proceeds on both sides of the statolith. The dashed line indicates the grinding plane.

3.2.4 Counting

Growth increment counting, i.e. the statolith reading, is the final phase of the whole

procedure and, under the assumption that one growth increment is deposited during one day,it allows estimation of the individuals age in days.

The statolith reading consists of the enumeration of the growth increments visible in the

statolith microstructure. Starting from the first growth increment around the focus, either the

dark or the light layers have to be counted, i.e. only the dark (suggested) or the light rings

(Figure 42). However, the natal ring has to be identified first in order to begin the counting

process.

In addition, since the increment width and visibility generally vary between the different

zones within the statolith microstructure, the best areas for increments identification and

counting also have to be identified.


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Figure 42. Ring identification for counting: either the dark or the light layers have to be counted. The arrows

point to the dark layers.

The natal ring is identifiable as a marked and complete check in the region immediately

around the focus. Its maximum diameter varies among species (from less then 100 to 250-

300 microns, Villanueva, 2000a,b; Steer et al., 2003; Sakai et al., 2004) and the check itself

can be more or less evident depending on the extent of the grinding process as well as on the

characteristics of the statolith (Figure 43).

Sometimes in the region immediately around the focus several checks are visible, making it

difficult to identify the proper natal ring. As general rule, the innermost complete marked

check should be considered as the proper natal ring4

(Arkhipkin, 1991; Jackson, 1994;

Arkhipkin and Perez, 1998; Villanueva, 2000a,b; Steer et al., 2003; Arkhipkin, 2004;

Ceriola, 2007).

Once the natal ring has been identified, the best areas (or best paths) to be followed for ringcounting have to be searched for in the statoliths microstructure. Statolith shape and

microstructure are species-specific so that growth increment identification can be easier in

some regions of the statolith rather than in others depending on the species. Moreover,

different regions of the statolith can be suitable for reading depending on the extent of

grinding. However, as a general feature, the narrowest increments are observed close to the

nucleus (in the postnuclear zone), while the broadest and more regular rings are observed

in the dark and peripheral zones.

The dorsal and lateral domes (in transversal or frontal section) have been shown to be the

best regions for increment counting (e.g. Jackson, 1989; Arkhipkin, 1991; Jackson and

Choat, 1992; Dawe and Beck, 1997; Bettencourt and Guerra, 2001; Quetglas and Morales-

Nin, 2004; Challier et al., 2005; Hendrickson and Hart, 2006; Ceriola, 2007; Jackson et al.,2007), but in some cases (mainly in loliginids) the rostrum in transversal section has been

preferred (e.g. Natsukari and Komine, 1992; Bettencourt et al., 1996; Arkhipkin, 1997; Raya

et al., 1999; Arkhipkin, 2005). Accordingly, it is within these regions that the best path for

counting is to be sought (Figure 44).

4 As already mentioned (see page 22), generally in the ommastrephids the natal ring is the first ring after

hatching and can be identified in the microstructure as the innermost ring, while in many loliginids several rings

are deposited during the embryonic development but usually not as marked/evident as the natal ring, so for

these species the innermost marked ring should be considered the natal ring. Ultimately, only individual

experience will help identify the proper natal ring both in species with embryonic increments and in

individuals with several checks in the postnuclear zone.

1

2

4

5

3


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Figure 43. Natal rings in statolith microstructure of adult specimens.

Natal ring

Natal ring

Natal ring


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Figure 44. The best path for ring counting is indicated by progressive numbers. WA = White Area; AA =

Adjacent Area where the number of rings of the WA can be estimated from.

3.2.4.1 Direct counting

Place the microscope slide with the ground statolith under the microscope and identify the

best magnification for ring identification; find the natal ring and the region/s where the

1

2

3

WA

AA


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increments are most visible; try to define a path to follow from the natal ring to the statolith

edge and begin counting, with the natal ring as increment number one.

Due to the plane of grinding, ring definition may change from the innermost to the external

regions. In this case it is necessary to slightly change the focus (by gently focusing) toachieve the best resolution; in this phase, the use of polarized light may help.

When counting in different regions (e.g. along the lateral dome for the postnuclear zone, off

the dorsal dome in the dark zone and in the lateral dome again for the peripheral zone), it is

possible to move from one region to the otherby following the rings border (Figure 45).

Figure 45. Ring enumeration following a well defined path: begins from the right, continues on the left, goes

back to the right. Numbers are an exemplification.

If areas where rings are not visible (i.e. white areas) are present in the observed section, the

rings number in these areas can be approximated as the number of increments in the adjacent

(just above or below) region of the same width, since in adjacent regions ring width does not

change dramatically. However, this procedure has to be used with caution when white areas

occur in proximity of t