Atlantic Sturgeon Research Techniques

US DEPARTMENT OF COMMERCE 

National Oceanic and Atmospheric Administration 

National Marine Fisheries Service 

Northeast Fisheries Science Center 

Woods Hole, Massachusetts 

May 2010

NOAA Technical Memorandum NMFS-NE-215 
Recent Issues in This Series:

196. Essential Fish Habitat Source Document: Haddock, Melanogrammus aeglefinus, Life History and Habitat Characteristics. 

2nd ed. By Jon K.T. Brodziak. December 2005. vi + 64 p., 27 figs., 2 tables. NTIS Access. No. PB2006-103439. [Online publication 

only.]

197.Withdrawn from series. 

198. Essential Fish Habitat Source Document: Bluefish, Pomatomus saltatrix, Life History and Habitat Characteristics. 2nd ed. 

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199. Distribution and Abundance of Fish Eggs Collected during the GLOBEC Broad-Scale Georges Bank Surveys, 1995-1999. 

By John D. Sibunka, Donna L. Johnson, and Peter L. Berrien. August 2006. iv + 72 p., 28 figs., 1 table. NTIS Access. No. PB2008-

107379. [Online publication only.] 

200. Essential Fish Habitat Source Document: Black Sea Bass, Centropristis striata, Life History and Habitat Characteristics (2nd 

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201. U.S. Atlantic and Gulf of Mexico Marine Mammal Stock Assessments -- 2006. By Gordon T. Waring, Elizabeth Josephson, 

Carol P. Fairfield, and Katherine Maze-Foley, eds. Dana Belden, Timothy V.N. Cole, Lance P. Garrison, Keith D. Mullin, Christopher 

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202. Evaluation of Northern Right Whale Ship Strike Reduction Measures in the Great South Channel of Massachusetts. By RL 

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203. Essential fish habitat source document: Spiny dogfish, Squalus acanthias, life history and habitat characteristics, 2nd 

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204. An Evaluation of the Northeast Region’s Study Fleet pilot program and Electronic Logbook System: Phases I and II. By 

Michael C. Palmer, Susan E. Wigley, John J. Hoey, and Joan E. Palmer. December 2007. NTIS Access No PB2008-107374. 

205. U.S. Atlantic and Gulf of Mexico Marine Mammal Stock Assessments -- 2007. By GT Waring, E Josephson, CP Fairfield, and 

K Maze-Foley, Editors. November 2007. NTIS Access No _______. 

206. Growth of Black Sea Bass (Centropristis striata) in Recirculating Aquaculture Systems. By Dean M. Perry, David A. Nelson, 

Dylan H. Redman, Stephan Metzler, and Robin Katersky. October 2007. NTIS Access No. PB2008-107374. 

207. Analysis of Atlantic Sea Scallop (Placopecten magellanicus) Fishery Impacts on the North Atlantic Population of 

Loggerhead Sea Turtles (Caretta caretta). By Richard Merrick and Heather Haas. February 2008. NTIS Access No PB2008-107373. 

208. Global Applications of the Large Marine Ecosystem Concept 2007-2010. By Kenneth Sherman, Marie-Chirstine Aquarone, and 

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209. Impacts to Marine Fisheries Habitat from Nonfishing Activities in the Northeastern United States. By Michael R. Johnson, 

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210. U.S. Atlantic and Gulf of Mexico Marine Mammal Stock Assessments -- 2008. By GT Waring, E Josephson, CP Fairfield, and 

K Maze-Foley, Editors. February 2009. NTIS Access No _______. 

211. Trends in Selected Northeast Region Marine Industries. By Eric M. Thunberg. July 2008. NTIS Access No. 

212. Summary of harbor porpoise (Phocoena phocoena) bycatch and levels of compliance in the northeast and mid–Atlantic 

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D. Orphanides, and Melissa L. Warden. August 2008. NTIS Access No. 

213. U.S. Atlantic and Gulf of Mexico Marine Mammal Stock Assessments -- 2009. By Gordon T. Waring, Elizabeth Josephson, 

Katherine Maze-Foley, and Patricia E. Rosel. December 2009. NTIS Access No. 

214. Mortality and Serious Injury Determinations for Baleen Whale Stocks along the United States and Canadian Eastern 

Seaboards, 2004-2008. By Allison H. Glass, Timothy V.N. Cole, and Mendy Garron. May 2010.NTIS Access No.
Atlantic Sturgeon Research Techniques

Kimberly Damon-Randall1, Russell Bohl1, Stephania Bolden2, Dewayne Fox3, 

Christian Hager4

, Brian Hickson5, Eric Hilton4, Jerre Mohler6, Erika Robbins7, 

Tom Savoy8

, Albert Spells9

1

 National Marine Fisheries Service, Northeast Regional Office, 55 Great Republic Dr., Gloucester, MA 01930 2

 National Marine Fisheries Service, Southeast Regional Office, 263 13th Avenue South, St. Petersburg, FL 33701 3 Delaware State University, Dept. of Agriculture and Natural Resources, 1200 N. DuPont Hwy., Dover, DE 19901 4 Virginia Institute of Marine Science, College of William and Mary, P.O. Box 1346, Gloucester Point, VA 23062 5 US Fish and Wildlife Service, 5308 Spring St., Warm Springs, GA 31830 6 US Fish and Wildlife Service, P.O. Box 75, 308 Washington Ave., Lamar, PA 16848 7

 Atlantic States Marine Fisheries Commission, 1444 Eye St. NW, Washington, DC 20005 8

 Connecticut Department of Environmental Protection, 79 Elm St., Hartford, CT 06106 

9

 US Fish and Wildlife Service, 11110 Kimages Rd., Charles City, VA23030 

US DEPARTMENT OF COMMERCE 

Gary Locke, Secretary 

National Oceanic and Atmospheric Administration 

Dr. Jane Lubchenco, Administrator 

National Marine Fisheries Service 

Eric C. Schwaab, Assistant Administrator for Fisheries 

Northeast Fisheries Science Center 

Woods Hole, Massachusetts 

May 2010

NOAA Technical Memorandum NMFS-NE-215 

This series represents a secondary level of scientific publishing. All issues employ thorough internal 

scientific review; some issues employ external scientific review. Reviews are transparent collegial 

reviews, not anonymous peer reviews. All issues may be cited in formal scientific communications. 
Editorial Notes 

Information Quality Act Compliance: In accordance with section 515 of Public Law 106-554, the 

Northeast Fisheries Science Center completed both technical and policy reviews for this report. 

These predissemination reviews are on file at the NEFSC Editorial Office. 

Species Names: The NEFSC Editorial Office’s policy on the use of species names in all 

technical communications is generally to follow the American Fisheries Society’s lists of 

scientific and common names for fishes, mollusks, and decapod crustaceans and to follow the 

Society for Marine Mammalogy's guidance on scientific and common names for marine 

mammals. Exceptions to this policy occur when there are subsequent compelling revisions in the 

classifications of species, resulting in changes in the names of species. 

Statistical Terms: The NEFSC Editorial Office’s policy on the use of statistical terms in all 

technical communications is generally to follow the International Standards Organization’s 

handbook of statistical methods. 

Internet Availability: This issue of the NOAA Technical Memorandum NMFS-NE series is 

being as a Web document in HTML (and thus searchable) and PDF formats and can be 

accessed at: http://www.nefsc.noaa.gov/nefsc/publications/. 
iii

Table of Contents 

Background ..................................................................................................................................... 1

Introduction ..................................................................................................................................... 1

Identification and measurement ...................................................................................................... 3

Summary ...................................................................................................................................... 6

Training requirements .................................................................................................................. 7

Sampling Methodologies ................................................................................................................ 7

Egg and larval nets and mats/pads ............................................................................................... 7

Gill nets and trammel nets ........................................................................................................... 8

Trawls ........................................................................................................................................ 14

Electrofishing ............................................................................................................................. 14

Passive Methods......................................................................................................................... 15

Summary .................................................................................................................................... 15

Training requirements ................................................................................................................ 15

Resuscitation .............................................................................................................................. 15

Handling methodologies ............................................................................................................... 16

Short-term holding ..................................................................................................................... 16

Summary .................................................................................................................................... 19

Training requirements ................................................................................................................ 19

Anesthesia .................................................................................................................................. 19

Summary .................................................................................................................................... 22

Laparoscopy and related technologies, non-invasive procedures, and traditional methods of 

determining sex and stage of gonadal development ..................................................................... 22

Laparoscopy ............................................................................................................................... 22

Traditional Methods of Determining Sex and Stage of Gonadal Development of Atlantic 

Sturgeon ..................................................................................................................................... 26

Summary .................................................................................................................................... 29

Training requirements ................................................................................................................ 29

Gastric lavage................................................................................................................................ 30

Summary .................................................................................................................................... 33

Training requirements ................................................................................................................ 33

Tagging and marking .................................................................................................................... 33

PIT Tagging ............................................................................................................................... 33

Scute Marking ............................................................................................................................ 35

Coded Wire Tags ....................................................................................................................... 37

External Tags – (nontelemetry) ................................................................................................. 37

Biotelemetry ............................................................................................................................... 38

Summary .................................................................................................................................... 39

Training requirements ................................................................................................................ 40

Tissue sampling ............................................................................................................................ 40

Biopsy ........................................................................................................................................ 40

Blood sampling and caudal venous puncture............................................................................. 41

Genetic tissue samples ............................................................................................................... 42

Fin spines ................................................................................................................................... 43
iv

Salvage specimens ..................................................................................................................... 44

Contaminants sampling .............................................................................................................. 45

Summary .................................................................................................................................... 45

Training requirements ................................................................................................................ 46

Cryopreservation ........................................................................................................................... 46

Short term storage ...................................................................................................................... 47

Cryopreservation ........................................................................................................................ 47

Summary .................................................................................................................................... 47

Acknowledgements ....................................................................................................................... 48

References ..................................................................................................................................... 49

Appendix – Field Data Sheet ........................................................................................................ 63
v

List of Figures 

Figure 1. Ventral view of shortnose sturgeon (Acipenser brevirostrum; left; Museum of 

Comparative Zoology 54265, 435 mm FL, Connecticut River, MA) and Atlantic sturgeon 

(Acipenser oxyrinchus oxyrinchus; right; Virginia Institute of Marine Science uncataloged, 780 

mm FL, James River, VA); note short snout and wide mouth of the shortnose sturgeon. Scale bar 

= 2 cm. Photos: John Weinstein, Field Museum of Natural History (left); Eric Hilton, Virginia 

Institute of Marine Science (right). ................................................................................................. 3

Figure 2. Lateral view of shortnose sturgeon (Acipenser brevirostrum; left) and Atlantic 

sturgeon (Acipenser oxyrinchus oxyrinchus; right); note the small bony plates (scutes) 

highlighted by the black arrows above the anal fin in the Atlantic sturgeon (same specimens as in 

Fig. 1). Scale bar = 5 cm. Photos: John Weinstein, Field Museum of Natural History (left); Eric 

Hilton, Virginia Institute of Marine Science (right). ...................................................................... 4

Figure 3. Depiction of the five sturgeon measurements (drawing courtesy of Dr. Eric Hilton, 

Virginia Institute of Marine Science). ............................................................................................ 6

Figure 4. Atlantic sturgeon (Acipenser oxyrinchus oxyrinchus) size distributions taken in 

anchored gill nets in Virginia are illustrated above (Chris Hager, VA Sea Grant, Marine 

Extension Program, unpublished data). Data support claims by previous researchers that 6 in 

mesh is highly effective at collecting a wide size range of fish. It also suggests that larger mesh 

sizes result in greater size disparity. ............................................................................................. 10

Figure 5. A hooded stretcher (photo courtesy of Jerre Mohler, US FWS). ................................. 18

Figure 6. Atlantic sturgeon (Acipenser oxyrinchus oxyrinchus) in an immobilization stretcher 

(photo courtesy of Chris Hager, VA Sea Grant, Marine Extension Program). ............................ 18

Figure 7. Atlantic sturgeon (Acipenser oxyrinchus oxyrinchus) being anesthetized (Photo 

courtesy of Jerre Mohler, US FWS). ............................................................................................ 21

Figure 8. Laparoscopy. ................................................................................................................ 23

Figure 9. Insertion of cannula and insufflation technique. .......................................................... 25

Figure 10. A laparoscopy incision closure showing two single sutures. (Photo courtesy of Jerre 

Mohler, US FWS). ........................................................................................................................ 26

Figure 11. Gonadal tissue of immature sturgeons (Photos courtesy of Jerre Mohler, US FWS). 

(Top photo) Immature male. Turgid, smooth white strip of testicular tissue surrounded by 

orange colored adipose tissue. (Middle photo) Immature female. Pinkish, grooved ovarian 

tissue surrounded by orange colored adipose tissue. (Bottom photo) Dissected section of 

immature male (top) and female (bottom) gonad. ........................................................................ 27

Figure 12. Gastric lavage (South Carolina Department of Natural Resources). .......................... 30
vi

Figure 13. Generalized depiction of the gastric lavage technique for Atlantic sturgeon (drawing 

courtesy of Eric Hilton, Virginia Institute of Marine Science). .................................................... 32

Figure 14. Illustration of PIT tag location (indicated by white arrow; top), and photo of a 

juvenile Atlantic sturgeon (Acipenser oxyrinchus oxyrinchus) being injected with a PIT tag 

(bottom). Photos courtesy of James Henne, US FWS. ................................................................ 34

Figure 15. Scute removal (left), and marking and insertion of coded wire tags (right) in lake 

sturgeon (Acipenser fulvescens). ................................................................................................... 36

Figure 16. External tag attachment locations for Atlantic sturgeon (line drawing adapted from 

Figure 3; courtesy of Eric Hilton, Virginia Institute of Marine Science). .................................... 39

Figure 17. Caudal venous puncture (performed on a small, two year old fish). .......................... 42

Figure 18. Depiction of a pectoral fin spine (drawing courtesy of Eric Hilton, Virginia Institute 

of Marine Science). ....................................................................................................................... 44
BACKGROUND 

In May 2000, NOAA’s National Marine Fisheries Service (NOAA Fisheries Service) 

published “A Protocol for Use of Shortnose and Atlantic Sturgeon” (Moser et al. 2000) 

(hereafter, the Moser Protocol). This document provided the first guidelines for handling and 

sampling Atlantic Coast sturgeon and was designed to help standardize research methodologies 

for these unique fish. The Moser Protocol has proven to be effective and has been closely 

followed by many sturgeon researchers. In the period since the document was published, more 

emphasis has been placed on sturgeon research which has led to an increased amount of 

sampling. This increased sampling has provided much needed information about both shortnose 

(Acipenser brevirostrum) and Atlantic sturgeon (Acipenser oxyrinchus oxyrinchus), and also, a 

great deal has been learned about methodologies to reduce injury and mortality to the fish during 

research activities. As a result of the ongoing status review for shortnose sturgeon, it was 

determined that a separate protocol document for Atlantic sturgeon was appropriate. 

Consequently, a group was formed to revise the protocol to incorporate new technologies as well 

as revise some of the existing methodologies to further reduce the potential for injury and 

mortality to Atlantic sturgeon. If Atlantic sturgeon are listed under the Endangered Species Act 

(ESA), it is possible that NOAA Fisheries Service may use the techniques described in this 

document as a baseline for development of consistent sampling protocols. 

INTRODUCTION 

Atlantic sturgeon have been considered a Species of Concern since 1998 following the 

joint decision by NOAA Fisheries Service and the US Fish and Wildlife Service (FWS) that 

listing the species under the ESA was not warranted. Because of concerns over continued 

declining trends in some subpopulations, in 2005, NOAA Fisheries Service initiated a second 

review of the status of Atlantic sturgeon. A status review team (SRT) consisting of four NOAA 

Fisheries Service, four FWS, and three US Geological Survey (USGS) personnel participated in 

the status review process. The status review was examined and supplemented by eight state and 

regional experts who provided both their individual insights and additional information to ensure 

the report provided the best available data. Lastly, the report was peer reviewed by six experts 

from academia and received favorable reviews. In the status review report (72 FR 15865), the 

SRT concluded that Atlantic sturgeon in the United States should be divided into five distinct 

population segments (DPSs): (1) Gulf of Maine; (2) New York Bight; (3) Chesapeake Bay; (4) 

Carolina; and (5) South Atlantic. The SRT also recommended that three of the five DPSs be 

listed as threatened (New York Bight, Chesapeake Bay, and Carolina). The SRT determined that 

the remaining two DPSs had a moderate risk of becoming extinct, though there were insufficient 

data to allow for a full assessment of these subpopulations; thus, a listing recommendation was 

not provided. An additional finding of the SRT was the overall lack of basic biological data for 

many of the Atlantic sturgeon subpopulations. 

Based on the information in the status review report and other best available data, NOAA 

Fisheries Service is currently in the process of determining whether to list Atlantic sturgeon 

under the ESA, which could ultimately affect how and what type of research activities are 

conducted on this species. Given the current status of Atlantic sturgeon and the lack of data on 

many subpopulations, it is necessary to perform research activities in a manner that allows for 
2

crucial information to be obtained on Atlantic sturgeon subpopulations while minimizing 

potential adverse impacts on the species. The consistent methods and approaches across 

subpopulations as elucidated in the summaries of this document would benefit future 

assessments and ranking of comparative population health. Should Atlantic sturgeon be listed 

under the ESA, researchers should refer to any relevant regulatory documents and consult with 

NOAA Fisheries Service to determine if planned research (even research that follows the 

protocols herein) is permitted under the ESA, and whether there are special authorizations 

required or reporting requirements that must be satisfied. 

In order to provide information on how Atlantic sturgeon protocols should be developed, 

a workshop sponsored by NOAA Fisheries Service and the Atlantic States Marine Fisheries 

Commission (ASMFC) was held in November 2007. Workshop participants were asked to 

identify specific activities, techniques, and methodologies that should be included in an updated 

protocol document. Over 30 sturgeon experts from Maine to Florida attended this two day 

workshop, and a subgroup was formed to draft the document. These protocols have been 

developed by researchers who have many years of experience conducting these activities 

specifically on Atlantic sturgeon. 

The workshop participants agreed that the Moser Protocol represented a valuable 

resource for conducting research activities on both species and decided that it should be used as a 

template to develop the revised protocols for Atlantic sturgeon. In order to maintain one 

comprehensive document, this report incorporates new technologies along with some of the same 

information from the Moser Protocol. 

As indicated in the Moser Protocol, sturgeon present some unique challenges for 

development of standardized methods. North American Atlantic sturgeon occur in various 

coastal, estuarine, and riverine habitats along the Atlantic Coast from the Saint John River in 

Canada to the St. Johns River in Florida. The differences in habitat, both within and among river 

systems, and latitudinal differences in temperature and sturgeon life history have resulted in 

sampling methodologies that are often specific to a given region or time of year. Similar to the 

Moser Protocol, research methodologies for sturgeon from across their entire range of habitats 

and for all life stages that have been studied have been included. Specific research plans for 

Atlantic sturgeon should be developed by researchers based on the conditions under which 

research will take place and in accordance with these research protocols. Research techniques 

that are invasive and hold inherent risks to the well-being of Atlantic sturgeon should only be 

conducted by researchers with the appropriate level of training and experience. Guidance 

outlining appropriate means to gain sufficient experience has been provided immediately 

following specific discussions of each research technique in this document. Methodologies for 

culturing and long-term maintenance of Atlantic sturgeon in captivity have not been included as 

both are addressed in the Culture Manual for Atlantic Sturgeon (Mohler 2004). The authors have 

attempted to identify and address any new and emerging technologies in this document but 

recognize that technologies change and advance over time. Thus, this should be a living 

document that allows for new techniques to be incorporated as they prove to be successful for 

Atlantic sturgeon. These techniques should be reviewed and revised as necessary, approximately 

every three years. If new technologies or techniques emerge between review periods, interim 

revisions should be considered. 

 The objectives of this document are (1) provide Atlantic sturgeon researchers with 

guidelines for conducting research to ensure that the safest, most recent, and effective techniques 

are used; and (2) compile the most recent literature review to support the use of these techniques. 
3

IDENTIFICATION AND MEASUREMENT 

Two species of sturgeon are present along the east coast of North America: Atlantic 

sturgeon and shortnose sturgeon. The two species differ greatly in their maximum total length 

(TL:427 cm and TL:143 cm respectively; Dadswell et al. 1984; Bain 1997), but juvenile Atlantic 

sturgeon may easily be confused with juvenile and adult shortnose sturgeon because of overall 

similarity in their general body form. Thus, care must be taken for correct identification, 

particularly among small individuals. Snout length is not a reliable character for identifying 

these species. Despite its common name, the rostrum of shortnose sturgeon can vary 

substantially in its size and shape, from truncated and rounded to moderately long and sharply 

pointed, even in similarly sized individuals. This variation matches that found in other aspects of 

the anatomy of this species (Hilton and Bemis 1999). There is significant variation in the shape 

and length of the snout of Atlantic sturgeon at different life history stages (e.g., juvenile vs. 

adult), although this has yet to be quantified (Bain 1997; Eric Hilton, Virginia Institute of Marine 

Science, pers. comm.). The snout of Atlantic sturgeon generally is more sharply pointed than that 

of shortnose sturgeon (Dadswell et al. 1984; Figure 1), but again, the morphological variation 

(including ontogenetic and other allometric variation) has not been fully described or quantified 

in this species. 

Figure 1. Ventral view of shortnose sturgeon (Acipenser brevirostrum; left; Museum of 

Comparative Zoology 54265, 435 mm FL, Connecticut River, MA) and Atlantic sturgeon (Acipenser 

oxyrinchus oxyrinchus; right; Virginia Institute of Marine Science uncataloged, 780 mm FL, James 

River, VA); note short snout and wide mouth of the shortnose sturgeon. Scale bar = 2 cm. Photos: 

John Weinstein, Field Museum of Natural History (left); Eric Hilton, Virginia Institute of Marine 

Science (right). 
4

In addition to several internal and external osteological characters that might serve to 

distinguish juveniles and adults of the two species (e.g., pale vs. dark viscera in Atlantic vs. 

shortnose sturgeon, respectively, Vladykov and Greeley 1963; the shape of rostral canal bones, 

Hilton 2002; shape and arrangement of frontal bones and caudal lateral line scales, Eric Hilton, 

Virginia Institute of Marine Science, pers. comm.), several key external characters may be used 

to distinguish between the two species in the field. Juvenile Atlantic sturgeon have a solid 

darkly pigmented dorsum and light ventral surface, whereas juvenile shortnose sturgeon (less 

than c. 30 cm TL), although they have a dark dorsum and light ventral surface, also have dark, 

irregularly-shaped blotches along the length of their body. While Atlantic sturgeon usually have 

a series of bony plates in the region immediately proximal to the anal fin (i.e., between the anal 

fin and the series of lateral scutes), these plates have not been observed in shortnose sturgeon 

(Figure 2). These plates are generally larger than the irregularly shaped and randomly 

distributed bony elements found in the skin between the five major rows of scutes; however, they 

are smaller than the scutes and the bony plates found more posteriorly on the dorsal, lateral, and 

ventral surfaces of the caudal peduncle. 

Figure 2. Lateral view of shortnose sturgeon (Acipenser brevirostrum; left) and Atlantic sturgeon 

(Acipenser oxyrinchus oxyrinchus; right); note the small bony plates (scutes) highlighted by the 

black arrows above the anal fin in the Atlantic sturgeon (same specimens as in Fig. 1). Scale bar = 

5 cm. Photos: John Weinstein, Field Museum of Natural History (left); Eric Hilton, Virginia Institute 

of Marine Science (right).

The most widely used and seemingly most reliable character over a broad ontogenetic 

range that has been used to distinguish between the species is the ratio of mouth width to 

interorbital distance (Moser et al. 1998). Dadswell et al. (1984) reported that Atlantic sturgeon 

have a mouth width of less than 55% of the interorbital width (range 43-66%) and shortnose 

sturgeon have a mouth width of more than 62% of the interorbital width (range 63-81%). 

Unpublished data suggest that the range of variation may be greater for both species. Specimens 

from the Merrimack River (NH, MA) had the following measurements: Atlantic sturgeon, 44-

70%, mean 50%, n=14; shortnose sturgeon, 59-80%, mean 68%, n=11 (Micah Kieffer, USGS 

Conte Anadromous Fish Lab, unpublished data.). For fishes caught in Connecticut (Atlantic 

sturgeon are likely of mixed stock), the following measurements were collected: Atlantic 

sturgeon, 34-49%, mean 44%, n=67; shortnose sturgeon, 54-79%, mean 66%, n=80 (Tom Savoy, 
5

Connecticut DEP, unpublished data). From the Hudson River, the following measurements were 

taken for the two species: Atlantic sturgeon, 32-92%, mean 50%, n=442; shortnose sturgeon, 47-

96%, mean 75%, n=177 (Sweka et al. 2006) with the vast majority of Atlantic sturgeon 

measurements (440 of 442) between 32-76% (Jerre Mohler, US FWS, pers. comm.). The level 

of variation within these and other morphological characters have yet to be quantified for either 

species, either at a species-wide or population-to-population level. 

Bath et al. (1981) described differences between larvae and small juveniles (8.4-37.0 mm 

TL) of the two species based on specimens from the Hudson River (NY). They regarded the 

most reliable identification character to be the relative mouth width. Snyder (1988) corrected 

miscalculations from Bath et al. (1981) and concluded that ranges overlapped for larvae of the 

two species. Snyder (1988) also described differences between the larvae (post-egg resorption) 

of the two species that he considered more reliable than mouth width, such as the presence of 

melanophores on the ventral surface of the abdomen of Atlantic sturgeon (absent in shortnose 

sturgeon) and a shorter distance between lobes of the lower lip (less than 20% mouth width in 

Atlantic sturgeon vs. more than 25% in shortnose). Additionally, he found a difference of pelvic 

fin ray number (17-22 for shortnose vs. 26-33 for Atlantic) and anal fin ray number (18-24 

shortnose vs. 22-30 Atlantic) for specimens over 60 mm standard length. Scott and Crossman 

(1973) also reported differences in the dorsal fin ray (38-46 in Atlantic and 19-22 in shortnose) 

and anal fin ray (25-30 in Atlantic and 19-22 in shortnose) counts. However, Vladykov and 

Greeley (1963) suggested that dorsal and anal fin rays were too difficult to count accurately 

(because the fin rays are heavily branched and embedded in a thick layer of skin) and were 

therefore of limited value as a distinguishing characteristic. Vladykov and Greeley (1963) found 

a difference in the number of gill rakers on the outside of the first gill arch of specimens greater 

than 20 cm (17-27, average 21.6 in Atlantic sturgeon and 22-29, average 25.4 in shortnose 

sturgeon). 

Birstein et al. (1997) recommended a series of 14 body measurements and six meristic 

characters (scute counts) to be collected for all sturgeon species. Because there are only two 

species of east coast sturgeon, the authors have selected measurements that have proven to be 

effective at differentiating between Atlantic and shortnose sturgeon specimens. At a minimum, 

the five measurements specified below (Figure 3) should be taken for all individuals, particularly 

when there is a question of species identification. In order to ensure consistency among 

researchers for the various measurements, place fish on either their left or right side, and take the 

following measurements from that position. 

total length: straight line along the body axis from the tip of the snout to the tip of the tail 

(not following the curvature of the body) 

fork length: straight line along the body axis from the tip of the snout to the posterior 

edge of the fork of the tail (not following the curvature of the body) 

head length: straight line along the body axis from the tip of the snout to the posterior 

edge of the bone that forms the gill cover (i.e., excluding the soft opercular flap) 

interorbital width: distance between the lateral margins of the bony skull at the midpoint 

of the orbit 

mouth width: distance between the left and right inside corners of the mouth (i.e., 

excluding the lips); this should be measured with the mouth closed 
6

Figure 3. Depiction of the five sturgeon measurements (drawing courtesy of Dr. Eric Hilton, 

Virginia Institute of Marine Science). 

Mouth width and interorbital distance should be measured with calipers. Several 

published keys, recommendations, and reports have cited mouth width as including the lips (e.g., 

see figure in Birstein et al. 1997; described one way in the key but illustrated the other in Musick 

et al. 1994); following these measurements, the ratio of mouth width to interorbital distance will 

be slightly higher. In order to ensure consistency of measurements for specimens throughout the 

range of the species, record this measurement on the inside corner of the mouth (following 

Vladykov and Greeley 1963 and Dadswell et al. 1984; see Figure 3) as this is the most common 

current practice. Measuring fork length is recommended as a reference for individual size since 

total length is subject to a greater level of measurement error (e.g., in “pulling” the tip of the tail 

ventrally, thereby over estimating the measurement). Any obvious fin erosion that impacts the 

measurement should be noted. If the researcher is inexperienced or the fish is “questionable” 

(i.e., less than 1 m in length) and in areas where both Atlantic and shortnose sturgeon occur, 

mouth width and interorbital distance measurements are necessary to confirm identification. 

Although no one diagnostic characteristic can be used to distinguish between shortnose and 

Atlantic sturgeon, in combination, the above characteristics are appropriate.

Summary 

For proper species identification of juvenile fish, the following measurements should be taken on 

all individuals sampled: 

 Mouth width 

 Interorbital width 
7

In addition, for general information and consistency between studies, we recommend that the 

following measurements also be taken: 

 Total length 

 Fork length 

 Head length 

Training requirements 

Only relatively minor training is needed for identification and measurements to ensure 

accuracy of identification and consistency of measurements. Because of difficulty in 

distinguishing the two east coast species, particularly at small juvenile sizes, researchers should 

gain sufficient experience and familiarity with identification through examination of specimens 

of both species in consultation with experienced researchers. 

SAMPLING METHODOLOGIES 

Choice of sampling methods for sturgeon is influenced by targeted life stage, habitat, and 

water temperatures. Sturgeon are highly susceptible to capture in gill nets whether stationary, 

drifting or hung in a trammel net configuration (Buckley and Kynard 1985; Hoff et al. 1988; 

Dovel et al. 1992; Geoghegan et al. 1992; Kieffer and Kynard 1993; Moser and Ross 1995; 

Collins et al. 1996; ASSRT 2007; ASMFC 2007). Trawls can also be highly effective but are 

often inappropriate for estuarine or riverine use because of benthic and physical conditions (e.g., 

narrow passages and uneven, rocky bottoms). In regions where fyke and large hoop nets are 

used by commercial fishers, juvenile and subadult sturgeon are occasionally captured as bycatch. 

Because of their much larger dimensions, pound nets can accommodate and hold fish of 

considerable size. Recently, commercial pound nets in Canada have provided large samples of 

adult fish to local scientists, and sturgeon of lesser size are routinely captured in the Chesapeake 

Bay in such gear (Mike Dadswell, Acadia University, pers. comm.; Chris Hager, VA Sea Grant, 

Marine Extension Program, pers. comm.). It is important to point out that reward programs and 

cooperative sampling efforts with commercial fishers can be more cost effective than fishery 

independent research collection efforts and should be considered as a means to collect data. 

Baited trot lines have proved to be an effective method for collecting white sturgeon (Acipenser 

transmontanus) and may have potential for collecting Atlantic sturgeon (Elliott and 

Beamesderfer 1990). 

Egg and larval nets and mats/pads 

Collecting larvae and eggs requires specialized techniques and approaches. As with other 

fishes, eggs of several species of sturgeon have been successfully collected with egg mats 

(McCabe and Beckman 1990; Marchant and Shutters 1996; Sulak and Clugston 1998; Fox et al. 

2000). Egg mats can be used to collect eggs as they are deposited. Egg sampling pads (e.g., 

floor buffing pads, approximately 2 ft in diameter [Fox et al. 2000]) are only effective in the 

immediate vicinity of spawning as sturgeon eggs become adhesive following fertilization as a 

result of changes in the egg membrane which cause the egg to swell and become sticky (Mohler 

2004). Researchers who wish to remove fertilized eggs which are adhered to the sampling pad 
8

should avoid trying to pick them off, and instead cut the pads with scissors to prevent rupturing 

the egg. Individual pads should be removed from the water, quickly scanned for the presence of 

eggs, and if present, eggs should be counted before the pad is returned to the water at the site of 

collection to allow successful hatching. 

In other systems, eggs and larvae and early juveniles have been satisfactorily collected in 

cone (Kohlhorst 1976) or D-shaped nets (Taubert 1980; Kynard et al. 1999) and with the use of 

epibenthic sleds. Mesh sizes of 2 mm2

 trap sturgeon eggs and larvae while letting some debris 

pass through. The net is attached to a weighted and floated 1 m diameter steel ring that has been 

flattened to maximize contact with the substrate (D-shaped, Kynard et al. 1999). A 1 m square 

or 2 m x 1 m Neuston net can also be used. The net is attached to a Danforth or grapnel-type 

anchor via a short bridle. This arrangement allows the net to stand upright in currents up to 1.0 

m s

-1. Depending on the current velocity and amount of debris accumulation, such gear should 

be fished no longer than 10 min in areas of suspected spawning. A flow meter should be 

positioned in the mouth of the net to allow calculation of egg or larval densities per volume of 

water sieved. Such studies are best conducted with the aid of telemetry data from prespawning 

adults to identify likely spawning locations (Collins and Smith 1993; Kynard et al. 1999). Dshaped nets were used to capture eggs of Chinese sturgeon (Acipenser sinensis) in the Yangtze 

River for four years. Tens of thousands of eggs were captured when the nets were set in areas 

occupied by telemetered fish. Eggs were reared to juvenile stages and released into the river 

(Wei and Kynard 1996). 

Nets should be deployed beginning at the earliest time spawning would be expected. 

Nets should be equipped with velocity meters to allow the volume of water filtered to be 

estimated to develop an index of abundance and an estimate of spawning success (# ELS/ 

volume of water sampled) (Taubert 1980). Nets should be checked routinely. Because of the 

relative rarity of the species, discretion by NOAA Fisheries Service will be used to determine the 

number of eggs collected. 

Light traps have proved to be of little value for collecting Gulf (Acipenser oxyrinchus 

desotoi) or Atlantic sturgeon larvae. Small juveniles are rarely taken with traditional survey 

gear, although some success has been achieved with modified trawls (Hrabik et al. 2007; Doyle 

et al. 2008). The limited success of mobile sampling gear may, in part, be due to the apparent 

tendency of larvae and juveniles to seek out crevices in rough bottom across which seines and 

trawls have low collection efficiencies. Because of their preference for such niche providing 

habitats, habitat pots which contain protective structure may be an alternative approach for 

sampling. These pots may work best in systems where natural protective structure is limited. 

Gill nets and trammel nets 

Sturgeon are highly susceptible to gill nets; however, it is well established that gill net 

selectivity with regard to size and even species is a function of biotic and abiotic factors. Biotic 

factors including morphology, behavior, and vertical and horizontal distributions (Hamley 1975; 

Marais 1985; Reis and Pawson 1999; Machiels et al. 1994; Dickson 1989; Purbayanto et al. 

2000) and abiotic gear factors such as mesh size, twine material, twine diameter, hanging ratios, 

and tie downs influence species retention, size selectivity, and fishing power (Hamley 1975; 

Machiels et al. 1994; Hovgard and Lassen 2000; Yokota et al 2001; Holst et al. 2002; Gray et al 

2005; Hager 2007). The historic commercial fishery for Gulf sturgeon provided evidence of age 

specific differences in gear vulnerability with larger and smaller sturgeon escaping preferentially 

(Huff 1975). 
9

It is easy to conceptualize why gill nets that cover more of the water column, by being 

longer or taller, may increase interaction rates and result in higher catch per unit effort (CPUE). 

Other gear construction alterations also significantly affect the species specific retention rates of 

the gear. The assumption that fish are simply gilled (i.e., are prevented from backing out of the 

webbing by a mesh caught behind the gill plates) by gill nets is false. Fish and other animals are 

retained in the nets in numerous ways, and the degree to which a species is retained is influenced 

by its body size and configuration of the gear it encounters. Organisms can be wedged (i.e., held 

by a mesh or meshes around the body) or become entangled by unique morphological attributes 

such as teeth, maxillae, scutes, snout, or other projections (Hamley 1975). Entanglement often 

results in struggling that subsequently wraps the animal in additional webbing. Gill nets can also 

be constructed so that they contain pockets or funnels of loose mesh that simply entrap and hold 

species until harvest. Tie downs create such consecutive bags of webbing and are used to 

increase likelihood of entanglement and entrapment between mesh walls. Finally, a net can be 

fished in a specific manner that promotes retention. Shad fishermen using anchored gill nets 

traditionally fish on slack tide to promote retention. The species has reduced/weak opercle plates 

and thus, typically does not gill well. Net retrieval in slack water allows fishers to literally scoop 

the fish up in the mesh and prevent them from falling free. This confinement of fish between 

webbing walls also promotes sturgeon retention upon gear retrieval as the weight of the fish is 

supported across numerous meshes instead of being concentrated on only a few. 

Sturgeon are morphologically unique. Their cone shaped snout rapidly transfers meshes 

over the head and along the body and thus, they are rapidly gilled or wedged. Their unique 

dermis covered with bony scutes increases the likelihood of entanglement and wedging, as 

meshes are rapidly caught as the fish attempts to pass through or around webbing. Larger fish 

may subsequently become wrapped in webbing once entangled while struggling, and smaller fish 

may be held by a single monofilament strand hung around a scute. Considerable overlap 

between sturgeon size distributions retained in varied mesh sizes results from these highly varied 

forms of retention (Figure 4, Chris Hager, VA Sea Grant, Marine Extension Program, 

unpublished data). 

Abiotic factors also affect how a fish interacts with the gear, and thus, retention 

characteristics. Interaction trials on captive Atlantic sturgeon, which examined the effect of 

twine size, hanging ratio, and the use of tie downs on sturgeon retention, reveal that enlarging 

twine (0.4-0.52 mm), augmenting hanging ratio (0.5-0.625), and removing tie downs (30” tie 

downs on 45” net) all significantly reduce retention rates (retained/interacted) (Hager 2007). 

This finding indicates that gear alterations or deployment methods that increase stretch in 

webbing, mobility of webbing, or amount of webbing for a given area will increase the 

likelihood of retaining Atlantic sturgeon. Increased sturgeon size distributions have also been 

noted in drift gill nets (Moser et al. 2000). Such nets are not only characteristically loosely hung, 

but the lack of anchors or lead line results in fish or other animals easily becoming wrapped in 

the net; thereby, increasing the likelihood of retention. NOAA Fisheries Service’s (Northeast 

Region) observer data indicate an increase in the size distribution of sturgeon retained when tie 

downs are used in gill nets. Gill nets consisting of 30.5 cm (12”) stretch mesh tied down to 

121.9 cm (48”) retained fish from 17-250 cm in total length (ASMFC 2007, section by Chris 

Hager, VA Sea Grant, Marine Extension Program) with the average fish being 137 cm TL. This 

difference in size selectivity is likely due to the fact that tie downs increase webbing mobility 

and meshes per unit area and create bags of webbing that entrap fish. Such alterations likely 

increase retention rates of sturgeon given that they are easily entangled. 
10

Figure 4. Atlantic sturgeon (Acipenser oxyrinchus oxyrinchus) size distributions taken in 

anchored gill nets in Virginia are illustrated above (Chris Hager, VA Sea Grant, Marine Extension 

Program, unpublished data). Data support claims by previous researchers that 6 in mesh is 

highly effective at collecting a wide size range of fish. It also suggests that larger mesh sizes 

result in greater size disparity. 

Both monofilament and braided mesh gill nets have been found to be effective for 

capturing sturgeon, although they may have different retention characteristics. While fish are 

captured more efficiently with smaller diameter twine, sturgeon can easily break free of webbing 

that is weak. Lighter twine has also been known to cut into sturgeon and cause injury (Moser et 

al. 2000); therefore, when using larger mesh sizes, twine sizes should also be increased. When 

targeting adults with 15 cm (6”) stretched mesh, multifilament nets of at least 0.52-0.57 mm 

should be used; with larger mesh sizes of 25.4-35.6 cm (10-14”), twine sizes of at least 0.9 mm 

are preferred to prevent loss and/or injury. 

Trammel nets consist of overlapping gill net panels (usually three) of varied mesh sizes 

hung on a single top and bottom line. This configuration not only provides more meshes per unit 

area but increases the likelihood of gilling, wedging, and entanglement through the congruent 

application of varied mesh sizes. Trammels also entangle and entrap fish between their 

overlapping walls of webbing. Fish may penetrate the larger webbing of the outer wall but 

subsequently fail to pass through the smaller inner wall. Given this construction, it is obvious 

why trammel nets collect a wider size distribution of fishes than other gill nets (Moser et al. 

2000). Some researchers claim that the increased likelihood of entanglement that the gear 

affords may reduce chances of mortality, but no research has been done to substantiate this 

claim. 
11

Sturgeon are benthic feeders and are generally captured near the benthos unless they are 

actively migrating (McCleave et al. 1977; Moser and Ross 1995). During immigration into 

riverine systems, adult fish are often netted in the top half of anchored nets (Albert Spells, US 

FWS and Chris Hager, VA Sea Grant, Marine Extension Program, pers. comm.). This capture 

location within the nets suggests that sturgeon may use the higher flow speeds in the pelagic 

region to aid migrations. In general, stationary nets should be heavily weighted or staked and 

allowed to contact the bottom. Whenever possible, nets should be set perpendicular to the 

current. In areas of high velocity or with heavy debris loading, this is not feasible, and nets 

should be set in eddies, on the downstream side of islands, or parallel to the current in 

midchannel (Buckley and Kynard 1985; Kieffer and Kynard 1993; Moser and Ross 1993; 

Kynard et al. 1999). In many southern rivers, trammel nets are set during slack tide periods only 

to reduce stress on fish and debris loads. This deployment method also maximizes the gear’s 

inherent entanglement/entrapment methodology. 

Employing staked, floating gear or tie downs allows the fisher to control the depth of 

water fished. Staked gill nets are not anchored to the bottom but are attached at both the top and 

bottom line to a series of vertical stakes driven into the substrate. Stakes are placed every 15.2 m 

(50 ft) or so. In commercial fisheries, anchored gill nets of less than 91.4 m (300 ft) are rare, and 

offshore lengths can reach 914.4 m (3000 ft). Shortening intervals between anchor points 

reduces tension on meshes caused by hydrodynamic forces. This deployment methodology also 

helps reduce the chances of anchors becoming dislodged at either end or of the top and bottom 

line twisting upon themselves in the currents. High rates of mortality have been observed when 

a net is dislodged or becomes twisted around retained fish. 

Drift gill nets that contact the bottom can be highly effective if the bottom is relatively 

snag-free (O'Herron and Able 1990; McCord 1998). Drifting reduces debris loading because the 

nets move with the debris and thus, intercept less of it. Generally, the short soak times, reduced 

pressure on the webbing, and active fishing methods used in conjunction with driftnets also 

result in less injury to captured fish (Moser et al. 2000). In upriver runs and pools, very light 

leadline and large floats can be used. In tidal areas, buoyancy should be reduced and the net 

dragged along the bottom wherever possible (McCord 1998). 

Gill net sampling and routine handling under favorable conditions do not appear to cause 

undue harm to sturgeon. In temperatures not exceeding 17C, lake sturgeon (Acipenser 

fulvescens) captured in gill nets (24 h net sets) and Passive Integrated Transponder (PIT) tagged 

showed significant responses in several physiological stress indicators but recovered nearly 

completely within 3 days of sampling, with no documented mortality (Baker et al. 2008). In an 

effort to reduce stress during removal from meshes, the fish should be inverted and their eyes 

covered. In some cases, net meshes should be cut for rapid and safe removal; in particular, if the 

fish is gilled and/or meshes have entered the gill case. Every effort should be made to revive 

nonresponsive fish, as in many cases fish that appear to be dead have recovered. 

In some cases, significant sturgeon mortalities have been documented in gill nets and 

trammel nets (Kieffer and Kynard 1993; Moser and Ross 1993; Collins et al. 1996; Kynard et al. 

1999; Stein et al. 2004; ASSRT 2007). Mortalities were often associated with elevated water 

temperature, extended soak times (Miller 2007), and net interactions that prevented normal 

respiration (Albert Spells, US FWS and Chris Hager, VA Sea Grant, Marine Extension Program, 

pers. comm.) To decrease mortality risk, precautions should be taken to reduce fish stress from 

collection methodology and environmental and biological conditions. When air temperatures are 

below 0C, sturgeon should not be out of the water for more than a few minutes. Sampling fish 
12

in gill nets in elevated temperatures has also been shown to increase mortality (Murphy et al. 

1995) and protract recovery (Wilkie et al. 1996). All fish should be processed while underwater 

if possible, as tissues can freeze. 

Collins et al. (1996) observed significant increases in gill net related mortality in the 

commercial drift gill net fishery for southern American shad (Alosa sapidissima) when water 

temperatures exceeded 18°C. Based on a 5 year fisheries independent anchored gill net survey 

with standard 24 hour sets and numerous mesh sizes (4.88-12” stretched mesh, n=430, 6-24°C), 

mortality likelihood did not increase homogenously across mesh sizes with rising water 

temperature; at some mesh sizes, it was negatively correlated. Interestingly, predicted mortality 

significantly increased across mesh sizes with fish size, with larger adults experiencing higher 

mortality than smaller, more resident individuals. It is worth noting that adult fish were recent 

immigrants to the system and only common in elevated water temperatures (>18°C). These 

findings may suggest that osmoregulation alterations upon freshwater immigration and long 

migrations to spawning grounds are significant stressors for adults. Conversely, higher adult 

death rates may simply be a reflection of the fact that most of these adults were retained in large 

mesh nets after having been gilled in a manner that restricted their respiration (Chris Hager, VA 

Sea Grant, Marine Extension Program, unpublished data). 

Niklitchek’s (2001) bioenergetic analysis of juvenile Atlantic sturgeon suggests that 

temperature, dissolved oxygen (DO), and salinity all significantly affect metabolism and thus, 

stress. At normal DO (70% saturation), instantaneous daily growth for small juveniles of 

Hudson origin peaks at ~18°C. Above this temperature, growth declines from temperature stress 

alone. The growth curve shifts to the right (21°C) at higher DO (100%) and to the left (16°C) at 

lower DO (40%). Salinity alone can be an important bioenergetic limitation factor (Niklitschek 

and Secor 2005). DO and salinity are naturally augmented during summer in Atlantic estuaries; 

thus, this season may be critical with regard to juvenile habitat limitation. If habitat is being 

limited by rising water temperatures, physiological tolerances and thermal sanctuaries may 

become increasingly important to species recovery, as systems are likely to become more 

bioenergetically challenging as climate continues to warm. 

Atlantic sturgeon may encounter unfavorable environmental conditions in many estuarine 

systems (Niklitschek 2001). Niklitschek (2001) found that juvenile Atlantic sturgeon exhibited 

negative behavioral and bioenergetic responses (food consumption, routine metabolism) when 

water temperatures reached 28C. Recent studies have indicated that shortnose sturgeon 

acclimated to higher temperatures are more tolerant of elevated temperatures (Ziegeweid et al. 

2008a), and anecdotal field observations indicate possible latitudinal variation in thermal 

tolerance. Atlantic sturgeon from southern river systems have been safely captured in gill nets at 

temperatures exceeding 30°C (Doug Peterson, University of Georgia, pers. comm.), while in 

some northern rivers, visible stress symptoms have been observed when Atlantic sturgeon are 

sampled in somewhat lower temperatures (24°C; Gayle Zydlewski, University of Maine, pers. 

comm.). Tolerance to elevated temperatures and low dissolved oxygen concentrations also 

appears to increase with age (body size) (Ziegeweid et al. 2008b; Jenkins et al. 1993). 

 Further complicating identification of safe temperatures during which gill netting can be 

conducted are the additive and synergistic effects of dissolved oxygen and salinity on 

bioenergetic responses and survival of juvenile Atlantic sturgeon (Niklitschek 2001; Niklitschek 

and Secor 2009). A review of the relevant literature on the effects of hypoxic conditions on 

sturgeon species by Secor and Niklitshcek (2001) revealed that dissolved oxygen levels below 

3.3 mg/L, regardless of temperature, can cause mortality in both shortnose and Atlantic sturgeon 
13

juveniles. The authors further suggest that dissolved oxygen at 60% saturation (4.3-4.7 mg/L at 

22-27C) or higher is necessary for shortnose and Atlantic sturgeon to avoid bioenergetic 

responses. 

It is important to note that sturgeon physiological research has only been conducted on 

juvenile sturgeon of Hudson origin. Applicability of results to subadult and adult Atlantic 

sturgeon and/or fish from different DPSs is unclear. Adult anadromous fish often have very 

different metabolisms than do juveniles for obvious evolutionary reasons. In some species, 

latitudinal differences are so extreme that spawning adults die instead of converting their 

metabolic systems to handle the osmotic challenges associated with freshwater habitation. 

Genetic variation presumably imparts inherent advantages to a given population uniquely suited 

for the physical characteristics of its native habitat. Atlantic sturgeon DPSs are unique at least in 

part because of this assumed adaptation to each system of origin. Clearly, more research needs 

to be done to derive applicable physiological limitation models that sufficiently describe the 

diverse physiological tolerances within Atlantic sturgeon populations. Once appropriate research 

has been conducted, gill net soak time guidelines based on temperature and dissolved oxygen 

conditions should be identified to ensure that the effects of environmental conditions on sturgeon 

health are not compounded unnecessarily by sampling stress. In some cases, sampling outside of 

optimal conditions may be necessary. However, this is only appropriate for researchers who 

have extensive experience sampling in a given system, who have demonstrated the ability to 

adapt gill net sampling to minimize stress to sturgeons, and who have not observed elevated 

mortality when sampling under these conditions 

Though we are only beginning to understand how and to what degree retention can be 

manipulated through various gear configurations, it is increasingly obvious that gill nets vary 

with regard to their selectivity. Because of the characteristically large size variation of the 

species and varied selectivity from gill net configurations, caution should be taken if one is 

attempting to characterize size or age distributions of sturgeon with fishery-dependent gill net 

data or independent data when gear is not uniform in construction. Temporal and spatial aspects 

of deployment may be equally important (WE Pine and S Martell, unpublished data). 

Researchers have even suggested that fish avoid the gear once educated (Pine et al. 2006), 

though numerous repeat captures of a single Atlantic sturgeon on consecutive days and fish that 

have been repeatedly captured in the same region for several months would cast doubt on this 

theory as would controlled interaction experiments (Chris Hager, VA Sea Grant, Marine 

Extension Program, unpublished data). 

Differences in sampling methods between programs and even within programs must be 

taken into account even when comparing CPUE. For example, despite recognition of a 

relationship between area fished and a gear’s efficiency, CPUE is commonly expressed as catch 

per linear length which ignores the vertical component of the gear. Differences in data collection 

and analysis can easily influence model outputs to the degree that they are misguiding or 

severely limit their usefulness. The structure of the data derived in each program is driven by the 

sampling program’s collection methodology and biology of the species, which may be unique 

within a given system and is likely varied between DPSs. Power to detect alterations in 

populations is affected by the presence of larger populations, larger changes in population, a 

higher probability of capture, and longer sampling periods (Zehfuss 2000). To be able to 

estimate and time dependent capture probabilities, capture rates must be consistently relatively 

high for each age class during each sampling period in order to separate whether an animal was 

present and not caught or was not present (WE Pine and S Martell, unpublished data). Gear 
14

retention is obviously influenced by presence or absence of target species, but it is also 

influenced significantly by biotic and abiotic factors unique to each study. Inconsistency in 

sampling biases predicted capture probabilities and thus, mortality estimates. Though biotic 

differences will never be eliminated between systems, they can be minimized. Abiotic factors 

can be effectively addressed through standardization. 

Standardizing equipment and developing temporal and spatial consistency between 

monitoring programs would greatly benefit future population analysis and allow for comparative 

studies (WE Pine and S Martell, unpublished data). Such standardization between studies or the 

establishment of surveys within each DPS will be required to understand population alterations 

within a DPS, exchanges between DPSs, and to be able to establish research priorities under a 

DPS structure. 

Trawls 

Where benthic and hydrodynamic conditions allow the use of trawls, this gear can be 

effective for capturing sturgeon. Collins et al. (1996) found that 39% of all juvenile Atlantic 

sturgeon and 8% of the adult shortnose sturgeon tag-returns from fish tagged in the Altamaha 

River, GA, were from the commercial trawl fishery. Sampling of shortnose and Atlantic 

sturgeon was conducted in the tidal portion of the Hudson River, NY from 1975-80 using 604 m 

and 10.7 m semi-balloon otter trawl having mesh sizes of 1.3-6.5 cm (Dovel and Berggren 1983; 

Dovel et al. 1992.). Fish >200-mm total length were regularly caught, with most fish around 500 

mm. These trawls were fished for variable lengths of time (up to 50 min) at tow speeds of 4km/h 

(2.2 knots). The Connecticut Department of Environmental Protection’s Long Island Sound 

Trawl survey has captured over 400 Atlantic sturgeon, ranging from 625 to 2135 mm FL. 

Sturgeon have been collected every year since the study began in 1984 from all depth strata 

sampled (up to 30 m). Up to 60 Atlantic sturgeon have been collected in a single tow (CT DEP, 

unpublished data) with little obvious external damage. The Hudson River Utilities Monitoring 

Program has also conducted a standardized trawling survey since 1985 by using a 3 m beam 

trawl with 1.3- 3.8 cm mesh. This gear is towed for 5 min against the current, and adult 

shortnose sturgeon (500-100 mm fork length) are caught regularly. This sampling indicates that 

even a small trawl effectively captures sturgeon. NOAA Fisheries Service northeast observer 

data suggest that trawls have a significantly lower mortality incidence than gill nets (ASMFC 

2007). Nevertheless, gear and deployment variables (e.g., tow time, depth of fishing, retrieval 

speed) can and should be controlled by researchers to minimize the risk of mortality. 

Electrofishing 

Electrofishing has not proven to be an effective method of collecting sturgeon (Moser et 

al. 2000). The electric trawl developed by Aadland and Cook (1992) for the collection of benthic 

species in riverine environments may offer an improved electrofishing method. Studies to 

develop species specific electrofishing techniques should look to hatchery fish for research 

subjects. If efficient methodologies can be developed for sturgeon, electrofishing could possibly 

offer a valuable tool at least for collecting and monitoring juvenile sturgeon. 

If electrofishing is to be used in habitats that may contain Atlantic sturgeon at any life 

history stage, the lowest effective voltages should be selected to minimize impacts on Atlantic 

sturgeon, whether targeted or incidentally exposed during electrofishing operations. Holliman 

and Reynolds (2002) found that white sturgeon (24-54 cm) were at a higher risk for hemorrhage 
15

when exposed to 60-Hz pulsed direct current (PDC) as opposed to continuous DC; thus, use of 

PDC is discouraged in waters containing Atlantic sturgeon. Refer to Reynolds (1996) for 

guidance on selecting the intensity of the electric field to be employed. 

Passive Methods 

Advances in noninvasive marine system sampling methodologies such as sonar, video, 

and combinations of both are quickly making these methods a viable alternative to traditional, 

potentially mortality-causing methods. Thus, such noninvasive methods should be used when 

possible since none have been shown to negatively affect sturgeon behavior. Such methods can 

also be used in advance of traditional netting efforts to increase likelihood of success and reduce 

effort and potential for gear loss. 

Summary 

We recommend that demographic assessments through surveys using gill nets be 

established and standardized with regard to biotic and abiotic factors known to influence catch 

composition. Establishment and support for such surveys should follow the ASSRT (2007) 

suggestions in that populations that are highly reduced in abundance should be viewed as high 

priorities. Standardization of typical sampling gear containing numerous meshes is not enough. 

Gear should also be standardized with regard to temporal and spatial deployment and other 

abiotic factors that significantly affect retention rates. 

Training requirements 

Safety of researchers and of the sturgeon being collected is of paramount importance. 

When large fish are taken, the fish can do considerable damage to itself, gear, and collectors if 

not handled correctly and quickly (AFS 2004). In order to safely handle large Atlantic 

sturgeon, experienced individuals and a crew with predefined duties are recommended to 

minimize risk. Because of inherent risks to both fish and humans, all researchers who wish to 

conduct collection efforts are strongly encouraged to actively participate in collection efforts 

with an experienced researcher prior to conducting independent efforts. Sampling with 

noninvasive methods requires training only in order to effectively operate the equipment and 

understand the results. However, sampling with gill nets, trammel nets, and trawls should only 

be conducted by researchers with sufficient levels of experience, which will ideally be specific to 

a river system or region. 

Resuscitation 

At times, Atlantic sturgeon removed from fishing gear appear to be nonresponsive. It is 

often possible to resuscitate these fish by flushing water, preferably oxygen enhanced, over the 

gills until recovery is obvious from the fish’s desire to escape. The most effective way to 

resuscitate fish is through the mouth, as if the fish were swimming forward. Dragging the fish 

back and pushing it forward is not ideal, but if there is no other option, then the drag back should 

very gentle so that the gill filaments are not damaged (Joel Van Eenennaam, University of 

California, pers. comm.). Also, the amount of oxygen exchange provided with this retrograde 

flow while pulling back is less than that provided when moving the fish forward (Joel Van 

Eenennaam, University of California, pers. comm.). The forward motion should be a faster 
16

motion than pulling back. The best method is to have a pump and hose of freshwater directed 

into the mouth, and if the end of the hose is actually placed into the mouth, a soft piece of sponge 

should be used to keep the metal or hard plastic from injuring the inside of the fish’s mouth (Joel 

Van Eenennaam, University of California, pers. comm.). Holding fish in large floating pens or 

in onshore raceways has been proven to be a successful resuscitation technique for both Atlantic 

and shortnose sturgeon in the Hudson River, NY (David Secor, University of Maryland, pers. 

comm.). This procedure is most effective when carried out while the fish is inverted. A form of 

restraint may be necessary if fish size and conditions warrant caution. Resuscitation should be 

attempted on all nonresponsive fish until such time as it is determined to be ineffective. In some 

cases, nonresponsive fish have been tethered by the tail in local waters overnight and found in 

the morning to be recovered, so this process can be protracted. 

HANDLING METHODOLOGIES 

Short-term holding 

It is often necessary to hold sturgeon for short periods while fishing with nets, tagging, or 

collecting tissue samples. Various methods have been developed for holding captive sturgeon for 

processing depending on fish size and number caught. 

Net Pens: When water quality is acceptable, portable net pens are a good option for 

holding fish for processing. Mesh sizes should be large enough to allow for the free exchange of 

water but small enough to prevent entanglement of all sized fish being sampled. It is preferable 

to use nets with knotless webbing and have a means of covering captive fish to prevent sunburn. 

It is not recommended that fish be held for extended periods of time when water temperatures are 

high. Thermal maximums may be different for northern and southern populations based on their 

acclimation to local water temperature regimes and the rate of temperature change fish are 

exposed to during collection and holding (Ziegeweid 2006) (see discussion of thermal tolerances 

on page 12). 

Holding Tank: Holding tanks should be designed to accommodate the size of the fish 

being worked on and should follow guidelines provided in the Atlantic sturgeon culture manual 

(Mohler 2004). When fish are held on board a research vessel, they should be placed in tanks 

with a flow through water supply that allows for total replacement of the water volume every 15-

20 min or in static bath with oxygen supplementation. Static bath water should be exchanged to 

maintain water temperature (Mohler 2004) and quality. A sump pump can be equipped with a 

long (20ft) hose to allow collection from deeper waters. While total water volume in the tanks is 

not critical, it should be sufficient to entirely cover the fish and allow for adequate control of 

temperature and oxygen levels. Oxygen saturation must not exceed 110%; this can result in 

oxygen receptors on the gills sending a message to the brain to slow down respiration and 

thereby cause build up of CO2 (hypercapnia), which can be lethal (Molly Webb, Bozeman Fish 

Technology Center, pers. comm., 2003, in Golder Associates Ltd 2006, Crocker and Cech 1996). 

If pure oxygen cylinders are being used to augment ambient oxygen levels, it is important to 

have a means for frequently or constantly measuring “in tank” oxygen levels to minimize 

environmental stress. 

In static bath holding tanks, osmotic stress can be relieved somewhat by the addition of 

0.25-0.5% uniodized salt without anticaking agents (Brian Richardson, Maryland Department of 

Natural Resources, pers. comm.) for nonripe broodstock. Ripe, female Atlantic sturgeon 
17

broodstock should not be exposed to a salt bath because of unknown effects on egg maturation 

(Mohler 2004); however, other species such as white and green sturgeon (Acipenser medirostris) 

have shown no adverse effects (Joel Van Eenennaam, University of California, pers. comm). 

Tethering: Tethering should be a holding method of last resort as the possibility exists for 

removing the mucus and causing abrasions that can result in post handling fungal and bacterial 

infections (Brian Hickson, US FWS, pers. comm.) and has been associated with elevated stress 

(Dick et al. 2006). If tethering is used, it should be only as a short term method of holding, and 

provisions should be made to protect fish from exposure by locating them in areas protected 

from sun and cold. When tethering in a tidal zone, care should be taken to ensure lines are long 

enough to allow the fish to stay on the bottom. Tethered fish should never be pulled out of the 

water and hung by the tether for weighing or other purposes as this will result in tearing the 

mesenteries of large fish (David Secor, University of Maryland, pers. comm.). 

When multiple large fish are collected that cannot be safely held in a net pen or be moved 

into an onboard holding tank, then tethering may be an option to hold fish for later processing. 

Golder Associates LTD (2006a) describes the procedure used for tethering subadult white 

sturgeon: “This tether line (2.5 cm thick soft-lay cotton rope) is placed around the caudal 

peduncle of captured white sturgeon. A ‘hose noose’ (i.e., rope fed inside a garden hose to 

prevent rope-on-skin abrasion) can also be placed around the pectoral girdle, so that one 

crewmember can maneuver the fish, while another member lifts the tail and places the tether line 

around the caudal peduncle. The tether should be snug enough around the tail to prevent the fish 

from escaping, but not so tight as to cause abrasion.” 

To minimize potential damage to fish, fishermen participating in reward programs should 

be provided with training to safely handle fish and supplied with a portable net pen or dockside 

holding tank to minimize handling stress if the maximum window for holding captured fish 

continues to be 24 hrs (Brian Richardson, Maryland Department of Natural Resources, pers. 

comm.). 

Stretcher: A hooded stretcher is commonly employed to handle large fish (Figure 5). 

Guidance for construction of a stretcher for handling broodfish can be found in Conte et al. 

(1988). Generally, the fish is placed in the stretcher with the ventral surface facing up and head 

inserted into a hooded chamber that has a drain. For very large fish, the stretcher can be tethered 

to the side of the boat for collection of field samples. Smaller fish can be lifted on board and 

placed in a stretcher holder (Mohler 2004). Aerated water is pumped with an aquarium style 

sump pump from an appropriate source through a plastic tube into the sturgeon’s mouth. Flow is 

adjusted to a consistent outflow from both opercula. The stretcher holder should be designed 

with sufficient pitch to allow water to drain toward the hood. Fish should not be held in excess 

of 1 hour in the stretcher. Captured sturgeon often have extremely sharp scutes which can easily 

slice skin as well as stretcher material, thus it is recommended that gloves be used when handling 

sturgeon (Mohler 2004). 

Fish are often immobilized and calmed by being placed in the recumbent position 

(Mohler 2004). If further anesthesia is required, a recirculating aerated anesthesia solution can 

be prepared in a five gallon bucket placed under the stretcher drain. Anesthesia solution should 

be prepared and administered according to guidance supplied in this manual and should be 

appropriate for the procedures to be conducted (see section detailing laparoscopy procedure and 

section on anesthesia). 
18

Figure 5. A hooded stretcher (photo courtesy of Jerre Mohler, US FWS). 

US Navy immobilization evacuation stretchers have been successfully modified for 

sturgeon handling and surgical transmitter implantation (Chris Hager, VA Sea Grant, Marine 

Extension Program, pers. comm.) (Figure 6). These stretchers immobilize the sturgeon by 

physically restraining it in a stretcher containing rigid sides. Adjustable belts and an internal 

jacket minimize movement, and flaps allow access to the sturgeon for surgical procedures. This 

apparatus can also be sized with the addition of flexible foam to accommodate smaller fish. 

Figure 6. Atlantic sturgeon (Acipenser oxyrinchus oxyrinchus) in an immobilization stretcher 

(photo courtesy of Chris Hager, VA Sea Grant, Marine Extension Program). 
19

Prior to releasing a captured sturgeon, ensure that the swim bladder is deflated. The 

Moser Protocol states that “Sturgeon are physostomous and tend to inflate their swim bladder 

when stressed and in air. If this occurs, efforts should be made to return the fish to neutral 

buoyancy prior to or during release. This can often be achieved by propelling the fish rapidly 

downward during release. If the fish still has air in its bladder it will float and be susceptible to 

sunburn or bird attacks. Often the remaining air can be released by gently applying ventral 

pressure in a posterior to anterior direction.” Swim bladders can also be manually deflated with 

the techniques described in the laparoscopy section (below). 

When sampling for sturgeon downstream of dams, be aware that while the dams are 

generating, total gas pressure (TGP) of the water may be elevated. This condition can cause 

stress and in extreme cases gas bubble disease. When fish are retrieved from the depths in these 

conditions, mortality may result; therefore, researchers should test TGP percentage in locations 

and times when expected to exceed 100% and should exercise caution by scheduling sampling 

for times when TGP is low, especially if sampling for larval sturgeon. Counihan et al. (1998) 

found that larval white sturgeon exposed to 118% TGP developed gas bubble disease within 15 

minutes of exposure, but no mortality was associated with the disease during the 10 day 

exposure. At 130% TGP, survival at 13 days post hatch was only 50%. 

Summary 

For short term holding, we recommend maintaining fish in floating net pens or holding 

tanks with flow through water to maintain proper temperature and dissolved oxygen 

concentrations. Field researchers should always carry instrumentation for monitoring water 

quality when handling sturgeon and make efforts to acclimate fish when water quality conditions 

are sufficiently different between bottom and surface waters. Tethering fish should be the 

method of last resort for short term holding. 

Training requirements 

Training should consist at a minimum of reading the relevant handling materials included 

in this manual and working with an experienced researcher until comfortable with fish handling 

techniques. Participation in an AFS sturgeon field handling technique course could substitute for 

working with experienced researcher. Review of the Atlantic Sturgeon culture manual is 

recommended. 

Anesthesia 

Reasons for anesthetizing fish include: minimizing handling stress resulting from 

physical restraint, increasing safety for the fish and handlers, and reducing fish movement to 

facilitate performing a research-related procedure, thus providing for the greatest welfare during 

handling. However, whether fish feel pain is an ongoing debate, and the distinction between an 

anesthetic agent with analgesic properties and immobilizing drug is not always clear (Neiffer and 

Stamper 2009). We suggest at this time there is no one agent for fish which provides both 

analgesic and anesthetic properties which would prevent detection of noxious stimuli as well as 

block the physiological stress response to restraint. For procedures on Atlantic sturgeon which 

may require small incisions and can be performed quickly (<10 min) by experienced researchers, 
20

use of anesthesia may not be the best course of action to obtain the desired results. If used, 

anesthesia should be administered to provide the lowest stage and plane of anesthesia as possible 

to safely conduct the specific procedure, and for the shortest exposure time. For most 

procedures, this would be defined as the point at which there is loss of equilibrium and failure to 

respond to tactile stimuli. 

Some additional guidance is provided by the American Fisheries Society whose policy 

statement indicates that prolonged stressful restraint should be avoided, but in some cases, 

utilization of general anesthesia for restraint may be advisable. However, the benefits of 

anesthesia and potential effects on data derived from anesthetized fish should be weighed against 

results obtained from fishes that have not been anesthetized. The full range of potential effects 

on the subject fish, not just the anesthetic qualities, must be considered. Physiological stress 

from prolonged periods of restraint required for some procedures should be avoided if possible 

through the use of anesthesia (AFS 2004). 

With regard to Atlantic sturgeon, three sedatives were evaluated at 5°C and 15°C in 

Mohler (2004): (1) metomidate (current trade name: “Aquacalm” by Aquatic Life Sciences, Inc., 

Ferndale, WA); (2) MS-222 or tricainemethane sulfonate (trade name: “Finquel”® by Argent 

Chemical Laboratories, Redmond, WA); and (3) 5% clove oil /95% ethanol mix (active 

ingredient in clove oil is eugenol). Because isoeugenol, a constituent of clove oil, is thought to 

be carcinogenic, its use on food fish is prohibited (AADAP 2008), therefore clove oil as an 

anesthetic for Atlantic sturgeon is not recommended. In the above evaluation, the appropriate 

dosage for metomidate and MS-222 was defined as the concentration necessary to sedate fish in 

3-4 minutes (i.e., loss of equilibrium and lack of response to tactile stimuli) and to allow 

recovery in 3-4 minutes, with the exception of metomidate, which required longer recovery times 

regardless of concentration tested (Wade Jodun, US FWS, pers. comm. in Mohler 2004). In 

general, Atlantic sturgeon took longer to recover at colder water temperatures. Metomidatetreated fish required the lowest dosage (15 mg/L as opposed to 200 mg/L MS-222) but took the 

longest to recover (>700 seconds at either temperature). Atlantic sturgeon (0.5-1 kg) were 

separately exposed for 20 minutes to the given dosages at water temperatures with no mortality. 

For larger Atlantic sturgeon (6-7 kg), the same dosages apply, but recovery times can greatly 

exceed those for smaller individuals. 

For subadult and juvenile Atlantic sturgeon, a simple water bath can be used to 

administer the sedative. For sedation of individual fish or those too large to place into a water 

bath, the desired solution can be administered via a recirculation system. This is accomplished 

by placing the Atlantic sturgeon onto a stretcher assembly and delivering anesthetic solution to 

the fish via an electric pump and tubing (Figure 7). This stretcher assembly may be designed to 

allow the delivered anesthesia to drain back to a reservoir and be used continuously in a 

recirculation fashion. If recirculation of anesthesia is used, an individual not directly involved 

with the surgery should be assigned the task of constantly observing the fish for effects of the 

sedative so that overexposure does not occur. With any sedative, risk of lethal overexposure 

increases if gill movement stops for an extended period of time; therefore, it is prudent to switch 

the circulation system to deliver fresh water once respiration frequency slows considerably. It 

should be noted that under anesthesia, opercular movement can cease or be reduced to nearly 

imperceptible levels for Atlantic sturgeon (unlike many other fishes), and it is suggested to use 

other monitoring methods in combination with observing opercular movement (such as heart 

rate) to ensure the safety of anesthetized fish. For this reason, it is also important to follow the 

recommended anesthetic dosages. 
21

Figure 7. Atlantic sturgeon (Acipenser oxyrinchus oxyrinchus) being anesthetized (Photo 

courtesy of Jerre Mohler, US FWS). 

MS-222- The concentration of MS-222 suggested for surgical implantation of 

transmitters should be sufficient to anesthetize the fish and eliminate any observable response 

during the procedure while allowing for rapid postoperative recovery; thereby, minimizing 

holding time. For surgical implantation of internal tags and procedures requiring similar 

incisions, sutures, and holding time, a simple anesthetic bath containing ambient water and MS222 at 50-100 mg/L is recommended (Harms and Bakal 1994; and used on Gulf sturgeon 

(Acipenser oxyrinchus desotoi) by Fox et al. 2000). It is advisable to check the pH of the 

anesthesia bath and adjust as necessary with sodium bicarbonate since MS-222 is a 

hydrochloride and can acidify water when used in freshwater settings. 

For invasive procedures (e.g., laparoscopy) that require prolonged surgical phase 

anesthesia, the anesthetic protocol for MS-222 detailed in the laparoscopy section of this 

document is more appropriate. When it is necessary to place Atlantic sturgeon under prolonged 

anesthesia, it is recommended that procedures be conducted in a controlled laboratory setting. 

Metomidate- If the intent for anesthesia is to minimize the stress response (suppression 

of the hypothalamo-pituitary-interrenal (HPI) axis and related cortisol release), metomidate is the 

only commonly used drug that performs this function. However, it is a hypnotic and does not 

induce general anesthesia as is evident in muscle fasciculations (i.e., twitching of small nerve 

fiber bundles) generally observed at the site of an incision and is probably a poor analgesic 
22

(Neiffer and Stamper 2009). A concentration of 15 mg/L is recommended for Atlantic sturgeon 

with sedation occurring in about 8 min (Jerre Mohler, US. FWS, pers. comm.). As previously 

stated, recovery times for fish treated with metomidate are much longer than for those with MS222. If metomidate or any other nonapproved drug is to be used on Atlantic sturgeon released 

back into the wild, the national Investigative New Animal Drug (INAD) coordinator at the US 

FWS’s Bozeman Fish Technology Center must be contacted for coordination purposes. See 

(http://www.fws.gov/fisheries/aadap/contactstaff.htm). 

Other investigative anesthesia drugs- At this time, there are two INADs for experimental 

anesthesia which can be used by registering with the Aquatic Animal Drug Approval Partnership 

(AADAP) office (for contact information see web address above). These additional drugs are 

known as AQUI-SE® and BENZOAK®. Currently, there is no withdrawal period as long as 

fish are not susceptible to harvest within 72 hours of administration 

(http://www.fws.gov/fisheries/aadap). Their current use is investigative, and they have not been 

evaluated for use with Atlantic sturgeon. 

Electro-anesthesia- Low intensity electrical currents have been used as a means of 

physical anesthetization in fish and provide an alternative to chemical anesthetics. Researchers 

have used alternating current (AC), straight direct current (DC), and pulsed direct current (PDC) 

to administer electronarcosis, and much of it is based on the same principles applied during 

electrofishing (Hartley 1967; Walker et al. 1994; Henyey et al. 2002; Barton and Dwyer 1997). 

The recommended method of applying electronarcosis to sturgeon is described by Henyey et al. 

(2002) using DC. Benefits of electronarcosis include shorter induction and recovery time, low 

risk of mortality, no visible sublethal effects (swimming and feeding behavior appear normal, no 

grossly apparent burns; Henyey et al. (2002)), and currents needed are imperceptible to 

researchers handling the fish and water containing current. Given these benefits, the utility of 

electronarcosis for even invasive techniques might prove to exceed that of chemical anesthetics 

such as MS-222. At this time, however, uncertainty exists over how electornarcosis works, and 

whether it is a true anesthetic or simply immobilizes and relaxes fish during research procedures 

(Hartley 1967; Henyey et al. 2002). 

Summary 

Because the administration of some anesthetics results in additional stress to fish, some 

researchers contend that it is an unnecessary and possibly risky procedure. However, others 

assert that for invasive procedures, it is more humane for the fish and safer for the researcher to 

administer anesthetics. We recommend that whenever it is practical for the researcher to 

administer anesthetics at the proper dosage, anesthetics be used both to reduce the risk to the 

researcher conducting the activity and to ensure humane treatment of Atlantic sturgeon. 

LAPAROSCOPY AND RELATED TECHNOLOGIES, NONINVASIVE PROCEDURES, AND TRADITIONAL METHODS OF 

DETERMINING SEX AND STAGE OF GONADAL 

DEVELOPMENT 

Laparoscopy 
23

Laparoscopy is a valuable surgical technique for examining the internal anatomy in 

sturgeon and has gained popularity in recent years since it is less invasive and provides a 

superior view of internal anatomy than do more traditional biopsy techniques which often require 

large abdominal incisions. For Atlantic sturgeon, laparoscopic procedures are mostly used to 

determine sex and stage of gonadal development, but they can also be used to observe other 

anatomical features for reasons of fish health. Laparoscopy requires specialized surgical 

equipment and should only be performed by individuals who are experienced or have had 

adequate training in laparoscopic techniques and anesthesia application. An experienced 

individual can determine sex and stage of gonadal development in as little as 15 min per fish 

(including anesthesia but not recovery) if no unforeseen complications arise. A minimum of two 

people are necessary to perform laparoscopy on sturgeon. Mature broodstock are often 2 m in 

length and often require more than two people to be safely handled. 

Laparoscopic equipment is of medical quality, requires a source of 110 Volt power, and 

is most suitable for use in controlled conditions with protection from the elements (Figure 8). 

The basic components of the system include: 

a. A stretcher or surgery table equipped with an anesthesia delivery system large enough to 

accommodate the fish being examined 

b. A light source with a flexible fiber-optic cable 

c. A 6mm stainless steel, rigid telescope which attaches to the fiber-optic cable 

d. A hollow cannula which is just large enough in diameter to permit insertion of the telescope. 

The cannula is equipped with a cutting tip and exterior threads which allow it to be screwed 

through the abdominal body wall of the sturgeon. 

e. A small video camera attached to the eyepiece of the telescope 

f. An LCD monitor upon which the internal anatomy is displayed 

g. A small air pressure/vacuum pump with flexible air lines 

h. A Verres insufflation needle 

i. Surgical supplies to make and eventually close the small incision (scalpel, suture material, 

needle holder, forceps, scissors, betadine antiseptic). PDS monofilament sutures have been 

recommended for sturgeon (Mohler 2004; Matsche and Bakal 2008). 

Figure 8. Laparoscopy. 
24

The standard operating procedures and details for performing laparoscopy on Atlantic 

sturgeon found have been described (Mohler 2004; Matsche and Bakal 2008); both citations 

recommend the following steps: 

1. Prepare the area where the procedure will be done, and assemble the laparoscopy equipment. 

2. Prepare anesthesia. There are two options for anesthetizing the sturgeon in preparation for 

laparoscopy: (a) use of a tub or other container large enough to immerse the entire fish to be 

anesthetized or (b) use of a stretcher assembly outfitted with a pump to recirculate anesthesia 

across the gills of the subject fish. For large, mature fish, a stretcher assembly with 

recirculating anesthesia system is sometimes more desirable since a much smaller total 

volume of anesthesia is required. Regardless of which option is used, prepare an “induction” 

dose of MS-222 anesthesia at 250 mg/L which is buffered with baking soda at 500 mg/L. 

Note** Do not dry mix the MS-222 and baking soda for any length of time before its use as 

the effectiveness of the MS-222 may be diminished. Also prepare a separate “maintenance” 

dose of MS-222 at 87mg/L buffered with 175 mg/L baking soda in preparation for the 

laparoscopy procedure. 

3. Administer the induction dose of anesthesia. Once the sturgeon is sufficiently anesthetized 

(see anesthesia section), the fish should be placed on the operating platform/stretcher with the 

maintenance dose of anesthesia recirculated across the gills to keep the fish immobile during 

the procedure. Always make sure that the dissolved oxygen levels are maintained between 

8–15 mg/L in the anesthesia solutions. One assistant should be designated to monitor the 

vital signs (i.e., opercular movement and heart activity). Opercular movement will slow 

considerably under anesthesia, but if all movement ceases, the fish should be quickly 

prepared for removal from the operating table or stretcher and resuscitated by gently moving 

the fish back and forth in an upright position in a tank of untreated water to flush the gills 

until rhythmic respiration is resumed. 

4. Make a small incision about 6-7 mm in length into the abdominal wall (Matsche and Bakal 

2008). For sex determination, the fish should be ventral side up with the body tilted 

somewhat so that the gonad will fall away from the body wall slightly to facilitate the sex 

determination. Look for a location on the fish’s right side between the third and fifth scute 

anterior of the pelvic fins and offset and from the abdominal midline to make the incision. 

Tip -Look for a favorable surface location which has fewer visible inclusions of dermal bone 

so that it will be easier to make and close the incision. The incision does not have to be 

completely through the body wall but should be large and deep enough so the tip of the 

threaded cannula will begin screwing itself through the body wall with its cutting tip as it is 

inserted in the incision and twisted. 

5. Screw in the cannula. Tip -Once the cannula begins to screw in, a slight upward pressure on 

the cannula while twisting may prevent damage to the intestines or other organs as it breaks 

through the abdominal wall and into the body cavity. Experience will allow you to determine 

when the cannula has finished cutting through the body wall and enters the cavity as you will 

feel less resistance when the tip breaks through. Alternatively, the telescope can be inserted 
25

into the cannula during entry to visualize the progress (Matsche and Bakal 2008). 

6. Insert the telescope. As you slide the telescope down through the cannula, you should begin 

to see some internal anatomy. If the view is obstructed, you may not have screwed the 

cannula in far enough to break through the body wall. If you can see some internal anatomy 

but the view is cloudy, the lens of the telescope may be smudged and should be wiped off 

before proceeding. If the view is still obstructed, you may need to insufflate the body cavity 

and/or deflate the swim bladder to create more open space. 

7. Insufflation of the body cavity can be achieved by attaching a low pressure air line to the air 

fitting on the telescope and introducing atmospheric air at no more than 1 L/min. via a low 

pressure pump (Figure 9). Air pressure should be less than 14 mmHg to prevent embolism or 

gas bubbles in the vascular system, which will lead to rapid death of the sturgeon. Because 

of this possibility, insufflation should be used sparingly. 

 

Figure 9. Insertion of cannula and insufflation technique. 

8. Determination of sex. Gonads lie adjacent to each side of the body wall and are examined for 

condition and texture for sex assignment (Bruch et al. 2001; Mohler 2004). 

9. Closure of incision (Figure 10). Normally, a single suture can be used for closure of 

laparoscopy incisions or punctures made with the Veress needle. Necessary instruments and 

other details concerning incisions closure are found in Matsche and Bakal (2008) and Mohler 

(2004). 

10. Post-surgery recovery. Move the sturgeon to its culture tank or back into the natural water 

supply, and orient the fish upright with a gentle back and forth motion to flush water across 

the gills. Once the fish is able to maintain itself in an upright position, it can be released. 

Normal recovery time is around 20 minutes but will likely be longer at cooler temperatures 

(Mohler 2004). 
26

Figure 10. A laparoscopy incision closure showing two single sutures. (Photo courtesy of Jerre 

Mohler, US FWS.) 

Traditional Methods of Determining Sex and Stage of 

Gonadal Development of Atlantic Sturgeon 

In circumstances where laparoscopy equipment is not available, yet sex must be 

determined, additional options may include: (1) traditional surgical biopsy (coeliotomy); (2) 

expression of sexual products; (3) blood plasma analyses of steroid levels; (4) ultrasound 

imaging; (5) other techniques such as the use of a borescope, meristics, and the shape of the 

urogenital opening. Each procedure is briefly explained below: 

1. Coeliotomy- This technique requires an incision though the ventral abdominal wall large 

enough to view internal organs with the naked eye. This procedure is very invasive and 

should only be performed by individuals who have had adequate training in both surgery and 

sex identification (Figure 11). This procedure is limited to larger fish (10 kg or larger) with 

visually differentiated gonads. Insert a blunt probe into the exposed coelom for manipulation 

of the gonad into a position where the germinal tissue can be viewed (Mohler 2004). This 

can be a difficult procedure because the germinal portion of the gonad lies against the lateral 

abdominal wall. For fish large enough to be sexually mature (about 130-150 cm in Atlantic 

sturgeon) it is relatively easy to identify sex with this technique because of advanced gonad 

development. Nonetheless, it remains highly invasive with increased risk of fish mortality 

when performed by untrained individuals. Sex can be identified after differentiation of the 

ovary and testis (the age and size of when this occurs is approximately 9 kg; Jerre Mohler, 

US FWS, pers. comm.). To determine the stage of gonadal development of small fish, a 

tissue sample of the gonad can be preserved in 10% buffered formalin, sectioned, embedded 

in paraffin, and stained for histological evaluation (Van Eenennaam and Doroshov 1998). 
27

Figure 11. Gonadal tissue of immature sturgeons (Photos courtesy of Jerre Mohler, US FWS). 

(Top photo) Immature male. Turgid, smooth white strip of testicular tissue surrounded by orange 

colored adipose tissue. (Middle photo) Immature female. Pinkish, grooved ovarian tissue 

surrounded by orange colored adipose tissue. (Bottom photo) Dissected section of immature 

male (top) and female (bottom) gonad. 

2. Expression of sexual products- If mature, sexual products may be expressed from the genital 

opening of wild individuals captured during spawning migration, especially in males (Mohler 
28

2004). It is extremely rare, but possible, to obtain ovulated eggs from the genital opening of 

wild females. In order for this to be the case, the female would have been captured in the 

process of egg deposition. 

3. Blood plasma analyses of steroid levels- This procedure requires that a blood sample be taken 

from the caudal vein and sent to a laboratory for analysis of the blood plasma hormones 

estradiol and 11-ketotestosterone via a procedure known as radio-immuno assay (Webb et al. 

2002; Fiest et al. 2004; Wildhaber et al. 2006). Obviously, this procedure does not give 

immediate results for sex assignment but can be valuable for determining sex and level of 

maturity in hatchery fish or analysis of the reproductive condition of wild populations. 

Blood samples must be collected, processed, and taken for analysis the same day. A 

centrifuge and means of keeping samples chilled are also needed along with specific 

techniques to obtain and ship the samples (Matsche and Bakal 2008). (See blood sampling 

and caudal venous puncture section below for more detail). In a recent study with 12 

hatchery Atlantic sturgeon ranging from 9–17 kg, 42% of individuals were correctly 

classified as to gender by using radio-immuno assay as verified by histology while 75% were 

correctly classified using laparoscopy (Jerre Mohler, US Fish &Wildlife Service, pers. 

comm.). 

4. Ultrasound imaging – Advancements in technology have made it possible to use portable 

ultrasound imaging equipment for identifying the sex of sturgeon, as well as determining 

other aspects of gonad development. Ultrasound offers a quick (10 seconds to 2 minutes per 

individual after significant experience), noninvasive method for sex identification (Chebanov 

and Galich 2009). In this procedure, the transducer is held against the posterovental portion 

of the abdominal region of the sturgeon. Specific characteristics of male and female gonads 

allow the reader to determine which gonad is being examined (testis or ovary) and at what 

stage the gonad has developed. Ultrasound imaging has been shown to be effective for 

determining the sex of the stellate sturgeon (Acipenser stellatus) (Moghim et al. 2002) and 

numerous other Eurasian species of sturgeon (Mikhail Chebanov, pers. comm.; Chebanov 

and Galich, 2009). In a study with shovelnose sturgeon (Scaphirhynchus platorynchus), 

Colombo et al. (2004) concluded that ultrasound imaging was 86% accurate and could be 

applied to other species of Acipenseriformes. In another study of shovelnose sturgeon 

(Wildhaber at al. 2005), however, ultrasound was shown to be less effective than use of an 

endoscope and resulted in correctly determining gender only 68-70% of the time, while use 

of an endoscope through an abdominal incision resulted in correct sex determinations 92% of 

the time. Bryan et al. (2007) also used ultrasound to study aspects of the reproductive 

condition of shovelnose sturgeon and pallid sturgeon (Scaphirhynchus albus), such as egg 

diameter, fecundity, and gonad volume. Although it was found that there was significant 

measurement error that required correction (e.g., egg diameter was underestimated by 52%), 

it was determined that ultrasound “accurately measure[d] the gonad volume, but needed a 

correction to accurately measure egg diameter and fecundity” (Bryan et al. 2007: 418). 

Because of variation in the anatomy of different species (e.g., thickness of body wall because 

of overall size of individuals; relative thickness and density of bony plates in the skin 

between the scutes), it is likely that there are species-specific aspects to interpretation of 

ultrasound data. At this time, no ultrasound studies have been performed with Atlantic 

sturgeon. The quality and ease of interpretation of ultrasound images will be dependent on 
29

the condition of the individual being evaluated. For instance, it has been well established 

that adipose tissue disrupts the ultrasound and prevents a clear image from being obtained. It 

is therefore suggested that ultrasound is most effective for sex determination after 

overwintering and before feeding is initiated (or in hatchery conditions, after a period of 

starvation) and large amounts of fat tissue are laid down and the gonads can be clearly seen 

in the ultrasound image (Chebanov and Galich 2009). Although ultrasound equipment costs 

are relatively expensive at this time, it is a promising method for quickly determining the sex 

of an individual by means of a noninvasive method given sufficient training in interpretation 

of ultrasound images. 

5. Other – A borescope was used by Kynard and Kieffer (2002) to determine gender in 

shortnose sturgeon by viewing the gonads through the urogenital duct, and they were able to 

determine the oocyte maturation stage by gonad color. However, they found no 

characteristic of a male testis that enabled them to conclusively distinguish between males 

and immature females. Bryan et al. (2007) used a flexible endoscope with female shovelnose 

sturgeon to determine egg diameter and fecundity by viewing the gonads through the 

urogenital opening as well as a small incision through the abdominal wall. A number of 

other techniques, such as meristics and the shape of the urogenital opening, have been tried 

with other species but have not been sufficiently tested on Atlantic sturgeon for reliable use 

at this time. 

Summary 

 There is no single gender determination technique which can be recommended for all 

occasions. Numerous factors relative to an individual researcher’s particular study must be taken 

into account before selecting one or more of the above-described techniques. Important factors 

to consider should include but are not limited to: (1) suspected level of sexual maturity likely to 

be encountered; (2) skill and experience of the researchers; (3) speed with which results are 

desired; (4) availability of equipment; and (5) number of fish expected to require analysis. 

Analysis of gonadal tissue samples via histology is the most accurate form of sex 

determination since specific structures associated with either spermatogenesis or oogenesis can 

be verified microscopically (Van Eenennaam and Doroshov 1998). For a more immediate sex 

determination of Atlantic sturgeon, laparoscopy gives the best view of internal organs in live fish 

and allows the most accurate determination of sex in individual fish having visuallydifferentiated gonads (usually fish >10 kg ). Use of laparoscopy is best suited to a hatchery or 

laboratory setting, but with the proper power supply requirements and overhead protection from 

sun glare and adverse weather, laparoscopy equipment can be set up on a vessel or shore-based 

operation near the capture location. Use of coeliotomy to determine gender is not recommended 

for most occasions since the size of the incision must be large enough (> 40 mm) to spread open 

the body cavity using a surgical retractor. Unless the fish is a reproductively mature individual, 

the target tissue that must be viewed lies obscured against the abdominal wall and is difficult to 

manipulate for a satisfactory view without some magnification or other technologic aid. 

Improvements in the above techniques and technologies will likely continue over time. 

Training requirements 
30

Mortality of research fish is less likely if individuals who wish to perform techniques 

such as laparoscopy or coeliotomy receive hands-on training with an experienced instructor 

using live fish. Expression of sexual products requires only a short amount of instruction from 

an experienced individual, but since live, ripe fish are difficult to have available for training, 

preserved or dead specimens could be substituted. Taking blood samples for the purpose of 

possible sex determination also requires hands-on training with live fish but mostly for success in 

locating the proper area and technique for extraction of the sample rather than prevention of fish 

mortality. 

Ultrasound imaging and meristic evaluations only require training in the use of the 

equipment and techniques but for proper sex determination, some training would be required 

from an experienced individual present. It would likely be possible to use fresh-dead or 

preserved specimens of known sex for this training. 

GASTRIC LAVAGE 

A variety of techniques for nonlethal sampling of stomach contents have been developed, 

but gastric lavage is recommended (Figure 12). Gastric lavage is relatively cost effective, 

nonlabor intensive, and reasonably safe and effective (Seaburg 1957; Foster 1977; Meehan and 

Miller 1978; Light et al. 1983; Haley 1998; Savoy and Benway 2004; Wanner 2006; Savoy 

2007; Shuman and Peters 2007). Only two papers have noted negative effects. Brosse et al. 

(2002) reported a statistically significant higher weight loss of lavaged sturgeon (7.97%) over a 

control group (5.84%), but all of the fish held lost weight over the 60 day holding period, 

indicating the presence of an additional stressor and confounding the results of that study. While 

Sprague et al. (1993) reported that 4 of 12 fish pumped died within 1 week of lavage from 

apparent water pressure damage, they provided few details on the technique, their experiment, or 

holding process. 

Figure 12. Gastric lavage (South Carolina Department of Natural Resources). 
31

Anesthesia: Haley (1998) and Savoy and Benway (2004) reported success while utilizing 

anesthesia (MS-222) during gastric lavage, as it is thought to relax muscles in the sturgeon and 

aid insertion of the lavage apparatus. Although this automatically extends handling time, use of 

anesthesia is encouraged to minimize the risk of injury during the procedure. 

Required use of anesthetics for lavage is still a question for debate. Anesthetics are only 

discussed in this section as they pertain to the lavage process. Additional information on 

anesthetics and their uses/limitations can be found in the anesthesia section. The Moser Protocol 

and some sturgeon researchers utilize anesthetics to calm fish and ostensibly to aid in insertion of 

the lavage apparatus into the pharynx. Use of chemical anesthetics automatically extends 

handling time to allow for fish to succumb and to recover from the anesthesia and requires 

additional space in which to anesthetize and recover fish. Additional concerns of the utilization 

of chemical narcotics involve the specific water chemistry where work is being conducted and 

interactions with the desired chemical agent. Working with salinities greater than approximately 

two parts per thousand or elements/solids/chemicals in solution may render some anesthetics 

ineffective (see anesthesia section above for full discussion). Lastly, researchers must take care 

to follow proper handling and disposal of treated water. Some researchers have noted that 

rolling sturgeon (and other fish) onto their backs calms them down enough to allow handling. It 

must be noted that this is not an absolute, and some fish may react strongly. In addition, vboards or props should be used to keep fish inverted and well supported all along the fish’s 

length. Given the size and strength of Atlantic sturgeon, restraints are advised for fish over 1 m 

TL to prevent injury to both fish and researcher. Ultimately, it must be noted that current 

information available about use of anesthetics, topically applied or otherwise, is equivocal as to 

whether they aid in carrying out the lavage procedure. 

Tube diameter and flexibility: Relatively flexible small diameter tubing is an essential 

part of this procedure. A small diameter tube (on the order of 2.0 mm outside diameter) is to be 

used for the average to midsized sturgeon (75.0-150.0 cm FL) to be lavaged, and smaller 

diameter tubes for the correspondingly smaller, immature fish. The flexibility of the tubing is an 

aid to prevent forcing the tubing through the walls of the alimentary canal. Aquarium tubing and 

the like should not be used owing to their stiffness. Haley (1998) specifically recommended 

intramedic type tubing over others because of its ductile nature and small diameter. The leading 

edge of the tubing should have all sharp edges blunted through heating or other manual means. 

While the flexibility of intramedic tubing seems to protect sturgeon from injury, it can take 

several attempts to get the tubing into the esophagus instead of curling around and exiting the 

mouth or through the gills. While unproven, forcing water out of the tubing while inserting the 

tubing into the alimentary canal is intuitively effective and should assist in allowing the tubing to 

enter the canal and prevent puncturing the walls of the canal by the tube. This may be an 

alternative to the anesthetic, which Haley (1998) noted might be required to aid in relaxing the 

muscular gizzard region of the alimentary canal of sturgeon (Figure 13). Researchers must take 

all care to prevent forcing the tubing into the fish and thus, causing damage. Gently moving the 

tube in and out while pumping seems to enhance the effectiveness of regurgitation. Large 

diameter tubes have been utilized by researchers to aid in inserting the flexible small diameter 

tube down the esophagus; they have been used as a sleeve to assist in getting the highly flexible 

small diameter tube past the oral cavity. This technique works well for some researchers and 

better for some species. Atlantic sturgeon have relatively narrow mouth widths for their size, so 

it is unclear if the two tube technique is applicable. 
32

Water delivery device: A variety of water receptacles and means of forcing water into 

the stomachs have been employed: syringes, garden sprayers (approx 2.5 gallon), hand operated 

and electric pumps. Regardless of the water delivery device utilized, it should allow researchers 

the ability to limit the amount of pressure in forcing water into the fish given the fragile nature of 

internal organs. If high volume or pressure pumps are used, a flow/pressure restricting device is 

imperative, although it cannot be stated what the upper pressure/volume limit is at this time. 

Positive results have been noted for both continuous water flow and pulsed or interrupted flow. 

No specific requirements can be made at this time without further directed study, particular to 

Atlantic sturgeon. Additional studies also need to be made on the effects of internal water 

chemistry, i.e. changing the osmotic balance of fish after introducing volumes of water to 

internal organs where fluid/chemical uptake is possible. Only Shuman and Peters (2007) have 

examined water chemistry after exposing shovelnose sturgeon to pulsed gastric lavage, and they 

report no negative effects. Additional research specifically on Atlantic sturgeon specifically is 

needed. 

Figure 13. Generalized depiction of the gastric lavage technique for Atlantic sturgeon (drawing 

courtesy of Eric Hilton, Virginia Institute of Marine Science). 
33

Temperature restrictions: Few specific temperature restrictions have been noted in 

published reports, but this may be a function of the small volume of work performed to date and 

general lack of work at temperature extremes. Shuman and Peters (2007) suggest water 

temperatures above 30oC may be problematic. Conducting lavage under freezing weather 

conditions may present unique dangers to the fish. General guidelines discourage exposure of 

fish to air temperatures below freezing for more than a couple of minutes rendering lavage 

ineffective, as it would be difficult to collect stomach contents while keeping a sturgeon 

submerged in water. Highly stressed individuals (from temperature change, capture stress, or 

other means) should not be subjected to lavage techniques. 

Summary 

 Gastric lavage is the least injurious, nonlethal technique available for examination of 

Atlantic sturgeon stomach contents. However, lavage is considered an invasive procedure, and 

full care and attention are necessary to minimize the potential for negative effects resulting from 

the procedure. As such, the smallest size water tubing feasible should be used with additional 

trade-offs between flexibility and ability to insert the tubing beyond the oral cavity. It is 

generally desired to hold fish in an inverted position, with the head end lower. Tubing must be 

inserted gently while expressing water. Tubing should never be forced if resistance is noted. It 

may take several tries to successfully get the tube into and beyond the pharynx. Volume of water 

and water pressure to be flushed through the fish also needs to be monitored and regulated. 

Continuous monitoring of the fish is required, and if water/discharge from the anus is noted, the 

tubing should be extracted somewhat and/or water pressure or volume reduced. Additional study 

of both short and long term effects, specific to Atlantic sturgeon are recommended. 

Training requirements 

Gastric lavage is a relatively simple but moderately invasive technique, and at a 

minimum, observation of an experienced individual and a review of Haley (1998) should be 

completed prior to performing gastric lavage on live Atlantic sturgeon. Brosse et al. (2002) and 

Buddington and Christofferson (1985) can be consulted for rough anatomy of the digestive tract 

of sturgeon as it pertains to lavage. Gastric lavage is likely to only remove those prey items in 

the esophagus and stomach or material located anterior to the gizzard. Although, digestion rates 

of sturgeon are generally unknown (Buddington and Christofferson 1985), the prey contents 

flushed from the beginning of the alimentary canal should probably be considered as recently 

eaten. Some polychaetes retrieved from shortnose sturgeon were noted to still be alive upon 

examination in the lab, several hours after the lavage (Savoy and Benway 2004). 

TAGGING AND MARKING 

PIT Tagging 

A number of Atlantic sturgeon population studies use passive integrated transponder 

(PIT) tags to provide long term marks. These tags are injected into the musculature below the 

base of the dorsal fin and above the row of lateral scutes on the left side of the Atlantic sturgeon 

(Eyler et al. 2009), where sturgeon are believed to experience the least new muscle growth. It is 
34

recommended that the needles and PIT tags be disinfected in isopropyl alcohol or equivalent 

rapid acting disinfectant. After any alcohol sterilization, we recommend that the instruments be 

air dried or rinsed in a sterile saline solution, as alcohol can irritate and dehydrate tissue (Joel 

Van Eenennam, University of California, pers. comm.). Tags should be inserted antennae first in 

the injection needle after being checked for operation with a PIT tag reader. Atlantic sturgeon 

should be examined on the dorsal surface posterior to the desired PIT tag site to identify a 

location free of dermal scutes at the injection site. The needle should be pushed through the skin 

and into the dorsal musculature at approximately a 60 degree angle (Figure 14). After insertion 

into the musculature, the needle angle should be adjusted to close to parallel and pushed though 

to the target PIT tag site while injecting the tag. After withdrawing the needle, the tag should be 

scanned to check operation again and tag number recorded. Some researchers check tags in 

advance and place them in individual 1.5 ml microcentrifuge tubes with the PIT number labeled 

to save time in the field. Because of the previous lack of standardization in placement of PIT 

tags, we recommend that the entire dorsal surface of each fish be scanned with a PIT tag reader 

to ensure detection of fish tagged in other studies. Because of the long life span and large size 

attained, Atlantic sturgeon may grow around the PIT tag, making it difficult to get close enough 

to read the tag in later years. For this reason, full length (highest power) PIT tags should be used. 

Figure 14. Illustration of PIT tag location (indicated by white arrow; top), and photo of a juvenile 

Atlantic sturgeon (Acipenser oxyrinchus oxyrinchus) being injected with a PIT tag (bottom). 

Photos courtesy of James Henne, US FWS. 

PIT tags far out perform external tags which are known to experience high shedding 

rates. However, laboratory studies indicate that sturgeon smaller than 200 mm TL shed PIT tags 

at a rate of over 50%, because of the less developed nature of the musculature at this size (Moser 

et al. 2000). Recent studies at the Bears Bluff National Fish Hatchery on shortnose sturgeon 

found that fish with a good condition factor could be tagged by a skilled operator with short 

(11.5mm) and long tags (14mm) at a minimum size of 300 mm TL with good survival (97%) and 
35

tag retention (98%) at 180 days post implant. There were no significant differences between 

cranial and dorsal implant of tags on weight gain or condition factor. Fish smaller than 250 mm 

were tagged only in the cranial location by necessity; retention rates were high (97%) but 

survival lower (70%) (James Henne, US FWS, unpublished data). Because of these high 

observed mortality rates and desire for consistency in implant location, sturgeon should not be 

tagged in the cranial location, and until safe dorsal PIT tagging techniques are developed for 

sturgeon smaller than 300 mm, only Atlantic sturgeon larger than 300 mm should receive PIT 

tags. Sutures were not used to close the PIT tag insertion point during this study (James Henne, 

US FWS, pers. comm.). 

Fuller et al. (2008) provide guidance on the quality of currently available PIT tags and 

readers and offer recommendations on the most flexible systems that can be integrated into 

existing research efforts while providing a platform for standardizing PIT tagging programs for 

Atlantic sturgeon on the east coast. The results of this study were consulted to assess which PIT 

tags/readers should be recommended for distribution. To increase compatibility across the range 

of these species, the authors currently recommend the Destron TX1411 SST 134.2 kHz PIT tag 

and the AVID PT VIII, Destron FS 2001, and Destron PR EX tag readers. These readers can 

read multiple tags, but software must be used to convert the tag ID number read by the Destron 

PR EX. The FWS/Maryland Fishery Resources Office (MFRO) will collect data in the coastal 

tagging database and provide approved tags for distribution to researchers. 

Golder Associates, Ltd. (2006b) reported on preliminary trials with a remote, underwater 

PIT tag reader equipped with an external antenna that successfully energized and recorded PIT 

tag numbers at a location baited to attract juvenile white sturgeon. When refined, this tool may 

prove valuable for studying seasonal habitat use and movement. 

Scute Marking 

Scute removal is a technique used by west coast sturgeon researchers as a secondary 

permanent mark to document fish that have been PIT tagged and injected with oxytetracycline 

(OTC) for staining hard parts in ageing studies (Figure 15). Rien et al. (1994) validated the 

technique with white sturgeon in the Columbia River Basin. In a two year study, 99% of fish 

marked by scute removal were clearly distinguishable upon recapture. Trained observers 

correctly read the mark with 93% accuracy while lay individuals read the mark correctly with 

74% accuracy. In sturgeon smaller than 2 ft, the scute can be removed with a number twelve 

curved scalpel blade. The lateral scute should be removed by cutting from the posterior margin 

and continuing anteriorly. The blade is angled slightly upward so the cutting surface lies against 

the lower scute surface. When the scalpel reaches the front of the scute, the thumb is placed on 

the posterior face and the scalpel is pulled up to complete the scute removal. 

Scalpels should be disinfected and allowed to dry between fish. For sturgeon larger than 

500 mm, a sharp filet knife can be used for scute removal. If skin is cut to the subdermis layer, 

antiseptic ointment should be available to apply to the wound. Fish can be anesthetized with a 

light dosage of anesthesia (75-90 mg/L MS-222) to immobilize the fish for this procedure (see 

anesthesia section for further detail). Fish larger than 300mm can safely be maintained at this 

dosage for 20 minutes. Care needs to be taken to ensure adequate oxygenation and temperature 

control in both the anesthesia and recovery chambers. During white sturgeon juvenile relocation 

into John Day Reservoir, fish are intramuscularly or intraperitoneally injected with LiquamycinLP (OTC) at a rate of 0.2 ml/kg to impart a mark on the pectoral spine for age validation studies 

(Apperson and Anders 1991; R.L. & L. 1996; Golder Associates, Ltd. 2006b). To prevent the 
36

chemical breakdown of OTC, keep the bottles in a cool area, protect them from direct light, and 

note the expiration date. A secondary benefit of this activity is that the antibiotic treatment 

minimizes possible bacterial infections resulting from fish handling, including scute removal. In 

addition to OTC, US FWS has been granted an INAD permit exemption allowing the use of 

calcein “SE-MARK®” to mark hard parts of fish, including sturgeon1

. 

 

Figure 15. Scute removal (left), and marking and insertion of coded wire tags (right) in lake 

sturgeon (Acipenser fulvescens). 

The lake sturgeon reintroduction program occurring in the southeastern United States has 

been using scute marking to aid in the identification of year classes of hatchery reared fish. The 

fish are marked as subyearlings (152-350 mm). Some fishery managers recapturing these fish 

have observed that as the fish grow there is a tendency for migration and joining of scutes 

adjacent the site of scute removal, making interpretation difficult. Marking subyearlings may 

require removal of multiple scutes to obtain a clear mark or switching from the left to right 

lateral scute on a rotating cycle. We recommend that a standardized procedure for assigning 

marks be developed for Atlantic sturgeon if scute marking is implemented. Until this is done, 

scute marking should be used on a case by case basis with mark location being coordinated 

among researchers. As scutes are thought to play an important role as an antipredation device, 

especially in small Atlantic sturgeon, we recommend that long-term studies designed to assess 

the impact of scute removal on mortality rates be conducted. 

 

1

 Calcein is currently available for use on potential food fish such as sturgeon under an Investigative New Animal 

Drug (INAD) permit exemption administered by the US Fish & Wildlife Service. Researchers desiring to use 

calcein for fish marking must register with the Aquatic Animal Drug Approval Partnership (AADAP) office in 

Bozeman, MT www.fws.gov/fisheries/aadap/calcein.htm to obtain a study number and an understanding of 

requirements for experimental use of calcein. Requirements for use of calcein on Atlantic sturgeon may differ 

depending upon federal listing status of the Distinct Population Segment of interest. 
37

Coded Wire Tags 

Coded wire tags (CWT) have been tested in several species of sturgeon with mixed 

results. Isely and Fontenot (2000) took a novel approach to use CWT to mark shortnose 

sturgeon by injecting sequentially numbered tags along the pectoral spine to allow for multiple 

recoveries during age validation studies. Retention rates were 78% for this placement. In the 

same study, placement of tags in the dorsal fin base and pectoral fin base were 98% and 96% 

respectively. Collins et al. (1994) found highly variable retention rates for shallow placement (1-

2 mm) of CWT in the snout cartilage of shortnose sturgeon (70-100% over the course of 400 

days). Bordner et al. (1990) working with white sturgeon juveniles (150-260g) had 100% 

retention for CWT placement below the dorsal scute and deep injection (3-4 mm) into the snout 

cartilage. Shallow placement (1-2mm) resulted in a 60% retention rate over 6 months. MS-222 

was used for anesthesia at a dosage of 90mg/L with no adverse affect on growth or survival. 

CWT injections were conducted without the use of a head mold. Mohler (1994) had 100% 

retention of CWT when injected underneath the first dorsal scute. As a result of insufficient data 

regarding effects of inserting CWT in the snout region and because of the presence of multiple 

sensory systems, we recommend that CWT be injected underneath the first dorsal scute for 

Atlantic sturgeon. The standardization of one location will help researchers in the conduction of 

broad-scale studies. Peterson et al. (2000) used CWTs to study population dynamics and wild 

juvenile shortnose sturgeon recruitment in the Hudson River system. CWTs provide an 

inexpensive mark to mass mark hatchery fish for use in mark–recapture studies to provide 

evidence of decreased recruitment of juvenile wild fish to the system. A CWT was detected in a 

captured 15-yr-old Atlantic sturgeon released into the Hudson River as a fingerling but even with 

the proper detector, numerous attempts were made before a positive detection was obtained. The 

presence of a visual secondary mark (i.e., pelvic fin removal) indicated the fish had received a 

CWT prior to its release, causing the researchers to perform repeat attempts at tag detection 

(Jerre Mohler, US FWS, pers. comm.). CWTs have limited utility for marking wild fish in the 

field because of the expense and complexity of the equipment but may be valuable tools for 

specialized studies using juvenile hatchery reared sturgeon that are too small to mark with PIT 

tags. Field researchers working in regions where these studies are being conducted should have 

access to a wand to detect CWTs during routine surveys. Because CWT equipment is expensive 

and has not been widely employed by Atlantic sturgeon researchers, its use is not recommended 

for routine studies. 

External Tags – (nontelemetry) 

The use of an internal tag (e.g., PIT) is preferable for long term mark-recapture studies 

because of higher retention rates (Isley and Fontenot 2000). However, there are occasions when 

having an external tag allows for identification of Atlantic sturgeon that are unexpectedly 

encountered by harvesters, the general public, and fisheries professionals. A variety of external 

tag types (e.g., dart, Carlin, and t-bar) have been used in a numerous locations (base of dorsal fin, 

scute drilling, and pectoral/pelvic fins) with mixed levels of success (Huff 1975; Carr et al. 1996; 

Collins et al. 1996). Researchers are encouraged to standardize the placement of external tags as 

much as possible. If an external tag is desired, researchers should contact the US FWS Maryland 

Fisheries Resources Office (MFRO). These t-bar tags should be implanted in a standardized 
38

location – based on experience of the authors, it is recommended that T-bar tags be placed at the 

base of the left dorsal fin musculature (Figure 16). 

If an existing tag is found in an Atlantic sturgeon at the time of collection, the researcher 

should pull lightly on the existing tag to estimate the level of attachment. If replacement is 

warranted, a new tag should be affixed (at the base of the left dorsal fin musculature). All data 

from both PIT tag and t-bar tagged individuals should be provided to the USFWS Atlantic 

sturgeon tagging database at the conclusion of the sampling season for inclusion into the coastwide tagging database. 

Biotelemetry 

As a result of high cost and difficulty in collecting Atlantic sturgeon, researchers 

conducting telemetry studies are often concerned with long term attachment/implantation of 

transmitters. Recent advances in battery/transmitter design have lead to longevities exceeding 10 

years. This increase in the longevity of transmitters when coupled with the highly migratory 

nature of Atlantic sturgeon makes the issues of system compatibility and tag code 

collision/redundancy an important issue that researchers must consider when designing and 

implementing telemetry studies. Researchers interested in conducting multiyear studies are 

encouraged to utilize internal implantation of transmitters (Dovel and Berggren 1983; Moser and 

Ross 1995; Zehfuss et al. 1999; Fox et al. 2000) to increase the likelihood of transmitter 

retention/attachment. Both anecdotal and published results (Smith et al. 1990; Savoy 1991; 

Zehfuss et al. 1999; and Fox et al. 2000) have shown that attachment rates for external 

transmitters are lower than for internal transmitters. Advances in casting materials used for 

transmitters have reduced rejection rates, but researchers who are concerned with maximizing 

retention rates may use a biologically inert compound (e.g., Silastic® Dow Corning) which is 

thought to reduce tissue reactivity and expulsion rates (Boyd Kynard, USGS Conte Anadromous 

Fish Lab, pers. comm.). Another low cost solution may be found in the use of beeswax, although 

reactivity rates for it are not well understood (Helm and Tyus 1992). 

Although retention rates of externally attached radio and acoustic transmitters have been 

shown to be lower (Zehfuss et al. 1999), there are occasions where external attachment may be 

warranted. Where planned research activities are scheduled for less than 4 months, researchers 

should consider external attachment, as it requires a less invasive procedure. Additionally, the 

use of pop-off satellite transmitters and archival tags requires external attachment for proper 

functioning. A number of techniques have been used to attach external tags including 

attachment at the base of the dorsal fin (Erickson and Hightower 2007; Erickson et al. in review) 

and by drilling holes through a dorsal scute (Edwards et al. 2007). In both the dorsal fin and 

scute attachment methods, a backing plate coupled with a sheath around wires/cables should be 

used to minimize irritation and maximize retention times. 

Fisheries scientists using radio or hybrid (radio/acoustic) transmitters, which necessitate 

an antenna for signal transmission, should consider antenna placement. Signal transmission 

values are maximized by trailing antennas, but the rate of infection and irritation via the opening 

in the body wall are considered greater than with internal antenna (coiled and trailing) (Knights 

and Lasee 1996; Bauer et al. 2005). Researchers are urged to consider the need for signal 

transmission values when considering either external or trailing antennas. Trailing antennas 

should only be used in cases where signal transmission range must be maximized (i.e., deep 

rivers, aerial surveys, dam passage studies). When a trailing antenna is required, researchers 

should consider the placement of the transmitter and the angle of the antenna at the exit point in 
39

the body wall (Bauer et al 2005). Researchers should utilize a shallow/more oblique angle of 

antenna exit as opposed to the antenna exiting at a perpendicular angle. The use of the 

shallow/oblique antenna is thought to decrease irritation and subsequent infection and to promote 

healing rates. 

Surgical implantation of internal transmitters should only be conducted on Atlantic 

sturgeon that are in excellent condition at times when they are not stressed from 

temperature/dissolved oxygen extremes (see discussion on tolerance to high temperatures and 

low dissolved oxygen conditions; 12). Anesthesia should be administered according to guidance 

provided in the anesthesia section of these protocols (beginning on page 19). All reasonable 

means should be used to check fish for existing biotelemetry transmitters prior to surgery. If it is 

determined that a fish has a currently functioning internal transmitter, implantation of a new 

transmitter is not recommended. Possible transmitter detection methods include hydrophones, 

coded wire tag wands, and metal detectors. Surgical implantation of transmitters should be 

conducted with sterilized transmitters and equipment to help minimize post operative infection 

rates (Mohler 2004). Surgery protocols should follow published guidelines for the location, size, 

and closure of incisions as well as application of betadine ointment (Conte et al. 1988; Fox et al. 

2000; and Mohler 2004). The intent of the betadine ointment is to coat the sutures until the fish's 

mucous has a chance to coat them (Robert Bakal, US FWS, pers. comm.). The betadine has 

antibacterial properties but also antifungal properties, which may be even more important 

(Robert Bakal, US FWS, pers. comm.). Upon completion of transmitter insertion, telemetered 

Atlantic sturgeon should be released immediately after regaining equilibrium at or near the 

location of capture. In cases where fixed collection gear remains in the area, the researcher 

should release Atlantic sturgeon in a location selected to minimize the potential of recapturing 

recently tagged individuals. 

Figure 16. External tag attachment locations for Atlantic sturgeon (line drawing adapted from 

Figure 3; courtesy of Eric Hilton, Virginia Institute of Marine Science). 

Summary 

 All Atlantic sturgeon greater than 300 mm should be tagged with PIT tags. Approved 

PIT tags and external t-bar tags should be acquired from the US FWS/MFRO office and data 

should be housed in their coastal tagging database. Because of the long life span and wandering 

nature of Atlantic sturgeon, researchers should consider both the long term and short term 

implication of any tagging studies. Researchers likely to encounter Atlantic sturgeon in other 

sampling activities should be provided the means to both scan captured Atlantic sturgeon and 

implant PIT tags in individuals not previously tagged. Scute removal as a means of externally 

marking Atlantic sturgeon should not be undertaken until more detailed studies on the long term 

effects on the health of Atlantic sturgeon are conducted. If an external tag is needed, researchers 

should contact the US FWS/MFRO to receive standard t-bar tags to be placed at the base of the 
40

left dorsal fin musculature. Given the high costs associated with telemetry studies, we 

recommend that researchers give consideration to the issues of system compatibility and code 

collision when initiating a telemetry study on Atlantic sturgeon. Short term telemetry studies on 

Atlantic sturgeon should consider the use of external transmitters while multiyear studies should 

utilize internal implantation techniques to minimize transmitter loss. 

Training requirements 

Because of inherent needs of sturgeon handling, all researchers who wish to conduct 

biotelemetry studies should participate in either hands on training from an existing or approved 

sturgeon researcher or an approved course on the surgical/handling procedures for Atlantic 

sturgeon. 

TISSUE SAMPLING 

Tissue samples for genetic analysis should be collected during sampling from all Atlantic 

sturgeon which are confirmed to be first time captures and not recaptures (confirmed while in the 

field through the presence of external tags or PIT tags). Collection of tissue samples for 

chemical analyses or age and growth studies is also strongly encouraged. Descriptions of tissue 

sampling techniques are provided below. Full use should be made of mortalities and salvaged 

specimens in order to maximize the information derived. 

Biopsy 

Collection of samples of internal organ tissues (e.g., for biological contaminant analysis) 

often requires sacrifice of a fish, salvaged specimens, or incidental mortalities, and may be 

obtained through dissection during a necropsy. However, biopsies can be taken of some organ 

tissues (e.g., gills, gonads, muscle) through surgeries with minimal impact on a fish. Muscle 

biopsies can be taken from the thickest portion of the dorsal (epaxial) musculature, as described 

in the Moser Protocol. After making a small incision, a biopsy can be used to take the tissue 

sample, and the wound closed with two sutures. 

Gonad biopsies in particular have been used successfully to identify sex and stage of 

gonadal development for several species of sturgeon and are commonly employed, particularly 

in aquaculture facilities. As Chapman and Park (2005) noted, “Although invasive, gonad 

biopsies in sturgeon cause minor trauma and remain the most reliable method of identifying their 

sex and stage of sexual maturity, especially at an early age.” Procedures for gonad biopsies are 

described in detail in several publications (Chapman et al. 1996; Fox et al. 2000; Webb and 

Erickson 2007). In short, as small as an incision as possible (25-40 mm) is made on the ventral 

surface of the anesthetized fish with a sterile scalpel. Once the gonad is located, a 1 cm3

 sample 

can be taken with a pair of forceps and a scalpel or a sample can be taken with an Eppendorfer 

biopsy punch or similar biopsy tool. The gonad sample must contain germinal tissue in order to 

be meaningful. The germinal portion of the gonad is oriented facing the body wall making a 

proper sample difficult to obtain in some cases. This is especially true when working with 

immature fish where a nonlethal sample is desired. In immature fish the gonad is composed 

largely of yellowish-orange adipose tissue, and if the biopsy sample contains only adipose tissue 

it will not reveal sex. Once the proper sample has been taken, the incision is then closed with a 
41

suture as described below. Also see section: (Laparoscopy and related technologies, noninvasive 

procedures, and traditional methods of determining sex and stage of gonadal development). 

Chapman and Park (2005) favored using Vicryl suture for closing the wound following 

biopsies because it did not require removal and did not irritate the skin as much as some other 

suture material. A more quantitative approach was taken in a comparative study which 

examined the tissue reactivity in a teleost fish. The findings of Hurty et al. (2001) suggest that 

Vicryl or cat gut materials, while cheaper, were found to cause more tissue reaction in comparion 

to absorptive materials (e.g., Maxon and PDS). Samples should be preserved in buffered 10% 

formalin for histological analysis or frozen for contaminant analysis. An alternative technique 

for gonad biopsy via laparoscopic techniques is also described in Matsche and Bakal (2008). 

Blood sampling and caudal venous puncture 

AFS (2004) cites three methods suitable for extracting blood from fishes: heart puncture, 

venous puncture, and caudal bleeding. Because of their morphology, heart puncture is 

impractical for sturgeon (i.e., the heart is well protected by the bony shields of the pectoral 

girdle). The preferred method for bleeding is through caudal venous puncture (venipuncture). 

Blood can be collected by puncturing the caudal vein (located ventral to vertebral column in the 

midline of the caudal peduncle) with a hypodermic needle or vacutainer (Figure 17). Unlike 

typical bony fishes, sturgeons do not have bony haemal spines protecting the caudal vein. 

Instead, the caudal vein runs in a canal formed by short, block-like cartilaginous vertebral 

elements; these elements are thickest on their lateral sides and are thinner ventrally. In caudal 

venous puncture, the needle is inserted at a 45° angle until the cartilage of the vertebral column is 

contacted (Stoskopf 1993); in large fish the vessels will be completely surrounded by cartilage, 

and the needle will need to be pushed through the thin ventral cartilage to puncture the vessel; 

care should be taken to not push too deep and puncture the notochord. The needle should then be 

backed off slightly and blood can be drawn (Stoskopf 1993). Negative pressure should be 

created in the syringe by pulling up on the plunger while inserting the needle so that blood will 

be drawn when the vein is punctured. While the needle should be inserted at an angle, this may 

not be possible or effective for juvenile fish, in which the needle may need to enter perpendicular 

to the vein for blood to be extracted (Figure 17). Because of the anatomy of the vertebral 

column and the caudal peduncle of sturgeon, a ventral approach for caudal venous puncture is 

most commonly applied. Alam et al. (2000) used 10cc disposable syringes with 20 gauge 

needles to draw blood from juvenile Gulf sturgeon; smaller or larger gauge needles may be 

appropriate depending on the size of the individuals being sampled. Glass syringes should not be 

used for collection of blood because of increased chances for premature coagulation (Stoskopf 

1993). In the field, blood samples should be stored on ice. Blood samples should be transferred 

to sterile containers (e.g., heparinized vacutainers) and need to be stored in a freezer (Alam et al. 

2000 stored samples at 0°C until sent out for analysis). For long term storage, samples should be 

maintained at -20 to -80°C (Molly Web, Bozeman Fish Technology Center, pers. comm.). In 

their study of bioaccumulation of metals and organochlorine compounds in Kootenai River white 

sturgeon, Kruse and Scarnecchia (2002) centrifuged samples to separate the plasma from the 

blood cells and froze the plasma until shipped to labs for analysis. Collection of plasma samples 

requires collection of blood with heparinized syringes or immediate transfer to heparinized 

vacutainers before centrifugation. 
42

Figure 17. Caudal venous puncture (performed on a small, two year old fish). 

Genetic tissue samples 

For genetic analyses, a 1 cm2 fin clip from one of the pelvic fins from living sturgeon 

should to be taken and placed in a labeled vial with an o-ring cap containing 95% nondenatured 

ethyl alcohol (EtOH) for genetic analyses (the pelvic fin is regarded as least intrusive, 

particularly for small individuals). It is recommended that the tissue be stored in a refrigerator 

for the first 24-48 hours (Tim King, USGS, pers. comm.). If long-term storage is necessary, the 

tissue should be placed in a refrigerator or freezer after the initial steeping and stored at 4°C to 

−20°C (Tim King, USGS, pers. comm.). This will aid in preventing evaporation of the ethanol. 

There may be some utility for collection of disease status information from pectoral fin clips. 

This should be considered on an as needed basis, with the standard sampling location for genetic 

samples being the pelvic fin. Fin clips provide sufficient DNA for extraction and analysis, and 

they are viewed as minimally invasive to the animal (AFS 2004). Tissue samples collected from 

dead sturgeon can include fin clips, the barbel, deep white muscle tissue (recommended if the 

animal is not freshly dead), liver, heart, or the viscous fluid in the eyes. Note that it is not 

recommended to clip barbels from living sturgeon for tissue samples. 

A relatively new tissue collection method for genetic samples is the use of FTATM cards 

(Whatman Inc., Clifton, NJ). Sample collection involves simple contact of a tissue to a specially 

coated paper which lyses the cells, stabilizes the DNA, and allows for long term sample storage 

at room temperature. The use of FTA cards is widespread, and they have been used for 

forensics, bacteriology, and plant and animal genetics (Purvis et al. 2006; Mbogori et al. 2006). 

Livia et al. (2006) found FTA-collected genetic samples to be reliable for microsatellite and 

Restriction Fragment Length Polymorphism (RFLP) analysis derived from samples of mucus 

and buccal cells of northern pike (Esox lucius) and brown trout (Salmo trutta). Borisenko et al. 

(2008) found the use of FTA cards to be less effective in a mammal survey than alcohol or 

cryopreserved tissue samples. They concluded that the observed DNA degradation was caused 

by high humidity (e.g., tropical conditions), tissue type that was sampled (e.g., liver, because of 

high enzymatic activity, degrades quickly; Hanner et al. 2005), or oversampling (e.g., too much 

tissue was blotted on the paper and the DNA did not fix properly). However, because of ease of 

use (including tissue collection from live specimens, such as buccal swabs, mucous samples, as 

well as tissues sampled through biopsies; Livia et al. 2006), transport (e.g., they can be sent 

through the mail with no need for special shipping with dry ice or alcohol), and archiving of 
43

samples, FTA cards may be suitable for many applications involving tissue collection for genetic 

studies of sturgeon. 

When collecting tissues, researchers should also perform the five measurements 

described on page 5, as well as collect information such as sex (if known), date of collection, and 

location (i.e., a GPS coordinate) of each fish sampled. The tissue sample (or a subsample, if the 

tissue is to be used by the researcher), the corresponding collection data, and any tag numbers 

available should be delivered to the National Ocean Service (NOS) archive in South Carolina 

(attention Julie Carter, NOS Marine Forensics Branch, 219 Fort Johnson Road, Charleston, SC 

29412). Because genetic information (e.g., natal rivers, etc.) is important for a broad range of 

aspects of sturgeon biology and management, results will be provided to the investigator who 

originally submitted the tissue; the investigators will also be acknowledged for their efforts in 

any resulting publications or reports. 

Fin spines 

The pectoral fin spine is the preferred calcified structure for ageing sturgeon (Brennan 

and Cailliet 1989). Removal or partial removal of a pectoral fin spine is nondeleterious (cf. 

otoliths, for which sturgeon must be sacrificed), allows for mark-recapture (e.g., OTC or calcein 

[see footnote on page 36]), and there is a precedence for this technique in the literature and 

previous care/handling documents (Cuerrier 1951; Rossiter et al. 1995; for Atlantic sturgeon 

specifically Stevenson and Secor 1999; Moser et al. 2000). There are some disadvantages to 

using this structure for ageing, including that the fin spine is composed of metabolically active 

material and calcium in the spine can be reabsorbed. The fin spine is an exposed structure and is 

susceptible to damage. This method for ageing has only been partially validated; see Rien and 

Beamesderfer (1994), Rossiter et al. (1995), Stevenson and Secor (1999), Whiteman et al. 

(2004), and Secor and Woodland (2005) for discussion of pros and cons and information on 

precision and validation of annuli counts for sturgeon fin spines generally. Split annuli, false 

annuli, inclusions of additional rays in the spine, crowding of annuli in older fishes, and 

difficulty defining the margin are concerns in using fin spines for ageing (Whiteman et al. 2004). 

Paragramian and Beamesderfer (2003) suggested that age estimates based on fin spines may 

underestimate true ages in their study of Kootenai River white sturgeon. Also, for markrecapture studies using OTC or calcein, it needs to be noted that left and right fin spines are not 

mirror images (note: spines marked with OTC or calcein must be stored in a dark location to 

prevent mark degradation). However, even with these disadvantages and caveats, fin spines 

remain the most practical structure for collecting age estimates for sturgeon.

Fin spine removal should follow the protocol outlined by Collins and Smith (1996), in 

which the spine is cut near its base with a minihacksaw (or similar saw or cutting instrument, 

e.g., bolt cutters, wire cutter, hacksaw, coping saw, or knife) and the more distal part of the spine 

is carefully separated from the fin rays with a scalpel. A recommended less invasive alternative 

technique involves taking a 1 cm section of the fin spine (Rien and Beamesderfer 1994; Secor 

and Woodland 2005). In this procedure, two cuts are made on either side of the section to be 

removed. In either procedure, when cutting the sample one should minimize the distance from 

the articulation without compromising the joint function or cutting into the basal recess of the 

spine, which houses a portion of the internal skeletal supports for the fin (Figure 18). After 

removing the sample, be sure to disinfect the wound and allow the fish to recover before 

releasing the fish. 
44

When collecting samples for ageing, researchers should minimize the amount of time that 

the sturgeon is out of the water. If the fish is to be held between the time of capture and tissue 

collection, minimize stress by using a holding pen or tank. Researchers may choose to use 

restraint or anesthesia (e.g., slings, stretcher, MS-222) when collecting samples, although 

anesthesia may cause additional stress, particularly during the spawning season. 

Most pectoral fins that have had the fin spines removed for ageing are expected to heal if 

the physical structure of the fin is not damaged (Collins and Smith 1996). However, there is 

evidence that large adults do not recover well after their leading fin ray is removed (Mark 

Collins, South Carolina Department of Natural Resources, pers. comm.), and it is encouraged 

that spines are not removed from larger adults because of potential deleterious effects. Full 

spines (i.e., including the base of the spine) may be removed from dead specimens. 

Figure 18. Depiction of a pectoral fin spine (drawing courtesy of Eric Hilton, Virginia Institute of 

Marine Science). 

Salvage specimens 

Dead or salvaged specimens can be invaluable for a number of basic and applied aspects 

of sturgeon biology and conservation. In the effort to maximize the information and data that 

can be derived from dead sturgeon, it is recommended that, depending on the condition of 

specimen (i.e., level of decomposition), tissues be taken as soon as practical, including fin clips 

for genetic analyses, muscle tissue for contaminant analyses, and fin spines for age and growth 

analyses. Deep muscles (i.e., close to the vertebral column) can be usefully sampled from even 

partially decomposed fish, and even badly decomposed specimens may prove valuable for 

osteological and other morphological and comparative analyses. 

It is important to maintain salvaged specimens and their derivative tissues so that they are 

available for future researchers (e.g., for morphological analyses, as voucher specimens for 

genetic studies). Natural history collections have great intrinsic value as a resource for future 

generations of researchers, students, fisheries agency officials, and resource managers. 

However, collection data for museum holdings of Atlantic sturgeon specimens are generally 

incomplete, particularly for large individuals (Eric Hilton, Virginia Institute of Marine Science, 

pers. comm.). These specimens are, therefore, of decreased value for many studies. Minimum 
45

standard data that should be kept with specimens include date of collection or recovery, locality, 

and the names of the collectors. 

Contaminants sampling 

Several tissues, including (but not limited to) gonads, muscle, liver, gills, and blood may 

be sampled for tissue residue analyses (e.g., dioxins, furans, PCBs, organochlorine pesticides, 

trace elements) to gauge effects of bioaccumulation and environmental contaminants (see review 

of contaminants in Gulf Sturgeon by Berg 2006). Depending on the analyses to be employed, 

tissue samples can be placed in chemical-clean jars (trace elements and organics), wrapped in 

aluminum foil, and placed in a zip-loc bag (organics), or wrapped in plastic wrap and placed in a 

zip-loc bag (trace elements). If the sample will be homogenized and split into aliquots for 

separate trace element and organics labs, chemical clean jars are easiest. Specimens should be 

stored frozen until analysis. MacDonald et al. (1997) and Agusa et al. (2004) stored their frozen 

samples at -20°C whereas Alam et al. (2000) stored specimens at 0°C. For some analyses (e.g., 

EROD/CYP1A analysis based on liver and gill tissue or that of sex steroids - estradiol, 

testosterone - and vitellogenin from blood plasma), tissues must be stored immediately on liquid 

nitrogen or dry ice and samples must be kept at -80oC prior to analysis. Brundage (2003) 

reported on analysis of ICP metals, mercury, Target Compound List (TCL) semivolatile organic 

compounds, organochlorines, PCBs, PCDD/PCDF and substituted isomers, and percent lipids 

from a shortnose sturgeon killed during dredging; the specimen was frozen after recovery. The 

specimen was partially thawed to allow for dissection of samples, which were kept on dry ice for 

transportation to the lab for analysis.

Although tissues generally degrade after death, with some analyses requiring fresh 

specimens that are immediately fixed or frozen, many usable tissues may also be taken from 

salvaged sturgeons; whether the carcass is suitable for tissue residue analysis is a judgment call. 

If the carcass is not too decomposed, muscle, liver, and gonad tissues can be collected; muscle 

samples should be taken as deep as possible. 

As an example, an Atlantic sturgeon carcass salvaged from Wellfleet, MA, in 2007 had a 

full screen for contaminant analysis. Samples were taken from the muscle, liver, and gonad. The 

samples were analyzed for organochlorine pesticides (n=23 compounds), dioxins (n=7 

congeners), furans (n=10 congeners), PCB congeners (n=95), polybrominated diphenyl ethers 

(PBDEs, n=40 congeners), and metals (n=19 elements). The cost was approximately $2,100 per 

sample (three samples for one fish; total cost approximately $6,300). The time required to 

complete the analysis ranges between 6 months to a year (metals may take only 3 months). 

Summary  Tissue samples collected from live or dead specimens can provide a variety of useful 

information to researchers. Biopsies of internal organ tissues are best suited for salvage 

specimens. When conducting muscle biopsies on live fish, tissue collected should be 

from the thickest portion of the dorsal musculature. 

 Gonad biopsies of 1 cm3

 to determine sex and stage of gonadal development can be taken 

with a 25-40 mm incision; to be effective samples must include germinal tissue, which 

can be difficult to obtain, especially in immature fish. 

 It is recommended that blood samples be taken by ventral venipuncture of the caudal 

peduncle; the needle should be inserted at a 45 angle, but in small fish this may not be 
46

possible, and an insertion perpendicular to the vein may be most effective. Blood 

samples collected in the field should be stored on ice; if long-term storage is desired, 

samples should be kept in sterilized containers at -20 to -80C. Blood should not be 

collected in glass syringes unless specifically requested in the protocol. Syringes and 

vacutainers can be used interchangeably as long as the type of vacutainer meets the blood 

collection objective (i.e. serum, vs. plasma). 

 For genetic analysis, we recommend that 1 cm2 tissue samples be taken from the pelvic 

fins of all fish, refrigerated or placed on ice for the first 24 hours if possible, and placed 

in 95% nondenatured EtOH; if long-term storage is desired, samples can be stored after 

initial steeping at 4 to -20C. Collecting genetic samples requires the use of clean, sterile 

instruments for each fish. A specific field protocol for sterilizing instruments would 

involve scrubbing flesh off the instrument and rinsing in fresh water then placing 

instruments in a 10% Clorox bath to degrade residual DNA. Instruments can then be 

rinsed in distilled water and dried before use. Ratio of ethanol to tissue should be 

approximately 10:1 for storing the tissue. A subsample of the genetic tissue sample and 

the corresponding collection data should be delivered to the NOS archive in South 

Carolina (attention Julie Carter, NOS Marine Forensics Branch, 219 Fort Johnson Road, 

Charleston, SC 29412). 

 A 1 cm section can be taken from the pectoral fin spine for ageing (see Rien and 

Beamesderfer 1994, and Secor and Woodland 2005 for technique details) but should not 

be taken from larger adults because of evidence that these individuals do not recover well 

from the procedure. 

 All savage specimens should be reported to NOAA Fisheries Service’s salvage network 

so that specimens can be obtained and sampled if desired; sampling of salvaged fish 

should be conducted to maximize the information gained. 

 

Training requirements 

The precise degree to which researchers should be trained for taking tissue samples 

should be matched to the level of invasiveness of the protocol. Because tissue sampling often 

involves a variety of specialized techniques and protocols, particularly for invasive aspects of 

sampling (e.g., biopsies, fin spine sampling, blood sampling), a high level of training or 

experience should be acquired by researchers for most tissue sampling activities. For taking fin 

clips for genetic sampling, a moderate amount of experience should be achieved so as to not take 

an excessively large portion of the fin. Tissue sampling from salvaged specimens requires 

minimal training, but experience and knowledge of particular protocols for sample preservation 

should be attained prior to sampling. 

CRYOPRESERVATION 

Collection and storage of gametes may play an important role in sturgeon aquaculture 

and conservation biology. Most effort has been made for cryopreservation of spermatozoa, 

although recent advances on the cryopreservation of eggs of other fishes have been made (e.g., 

salmonids; Kobayashi et al. 2007); these and other protocols may be extended to sturgeon in the 

future. For collecting gametes, see Mohler (2004). 
47

Short term storage 

Sperm stored undiluted in refrigerators (on ice and given oxygen daily) has been shown 

to retain its fertilizing potential for up to 14 days for paddlefishes, but only 5-7 days for 

shortnose sturgeon and Gulf sturgeon (Park and Chapman 2005) and with significant decrease in 

sperm motility. Dorsey (2009) found that wild caught Atlantic sturgeon had higher quality 

sperm than captive individuals, perhaps because of environmental or diet stressors on captive 

individuals (captive samples did not survive after 21 days). For short term storage of sperm, Bill 

Wayman (US FWS Warm Springs Fish Technology Center, pers. comm.) recommended keeping 

fresh sperm viable by keeping it in oxygen filled sealable plastic bags or centrifuge tubes on ice 

(DiLauro et al. 1994). Containers should only be filled half way with sperm in order to provide 

enough volume for sufficient oxygenation, and multiple samples should be taken per fish in 

order to ensure that some are viable (Curry Woods, University of Maryland, pers. comm.). 

Samples should be resuspended daily by gentle inversion, and oxygen should be replenished 

daily. Dorsey (2009) found that sperm stored with oxygen was of higher quality (viability, 

motility, curvilinear velocity, and cell ATP level) than sperm stored in an oxygen free 

environment. Various extenders have been used for storage and cryopreservation of sturgeon 

sperm, including buffered sodium chloride (NaCl) or potassium chloride (KCl) and saccharose 

solutions diluted one to one sperm to extender (e.g., Billard et al. 2004). Horvath et al. (2005) 

used a modified Tsvetkova’s extender in a 1:1 ratio with sperm to cryopreserve sperm of 

shortnose sturgeon. Bill Wayman (USFWS Warm Springs Fish Technology Center, pers. 

comm.) used HBSS-S at 100 mOsm/kg as an extender at a ratio of 1 ml of sperm to 4 ml of 

extender. Park and Chapman (2005: table 2) provide a protocol for mixing an extender for shortterm storage of sturgeon sperm (up to 28 days at 4oC). Dorsey (2009) tested two extenders and 

found that the one described by Park and Chapman (2005) in the presence of oxygen was most 

effective and recommended dilution immediately upon collection to avoid contamination issues. 

Cryopreservation 

Sperm can be cryopreserved with dimethyl-sulfoxide (DMSO), dimethyl acetate (DMA), 

methanol, or ethylene glycol; different cryoprotectants show different results depending on the 

species of sturgeon involved. No equilibration time may be needed for sperm that is frozen 

immediately (Jähnichen et al. 1999) but generally is no more than 10-15 minutes. Sperm can be 

frozen by directly placing pellets on dry ice, in vials or straws placed in programmable freezers, 

or by suspending straws or vials in a rack 3-5 cm above liquid nitrogen (Billard et al. 2004). 

Sperm can be thawed quickly in warm water (e.g., for 9 sec in a 40o

C water bath; Bill 

Wayman, USFWS Warm Springs Fish Technology Center, pers. comm.; see also Jähnichen et al. 

1999; Billard et al. 2004). Activators may also include sodium, calcium, or saccharose, which 

may improve sperm motility or fertilizing capacity (Billard et al. 2004). Changes in both 

motility and acrosome structure of cryopreserved sperm affect the fertilization potential of the 

sperm, but these may be offset by use of excess sperm during fertilization (Jähnichen et al. 1999; 

Billard et al. 2004). Bill Wayman (USFWS Warm Springs Fish Technology Center, pers. 

comm.) reported 10-20% motility of cryopreserved Atlantic sturgeon sperm when thawed. 

Summary 

 To maximize the success and value of collected sperm, we recommend: 
48

 Diluting sperm immediately after collection with an appropriate extender (as described 

by Park and Chapman (2005) 

 Store sperm in the presence of oxygen, and resuspend the sperm dailyTraining 

requirements 

Because cryopreservation involves specialized protocols, a high level of training or 

experience should be acquired by researchers prior to initiation of activities. 

ACKNOWLEDGEMENTS 

The authors would like to acknowledge the following people who provided helpful comments 

and suggestions in their reviews of this document: Dr. Mary Moser, Dr. Joel Van Eenennaam, 

and Dr. David Secor. 
49

REFERENCES 

AADAP (Aquatic Animal Drug Approval Partnership Program). 2008. AQUI-S use on food fish 

under IDAD 10-541 strictly prohibited. AADAP Newsletter 4-2:1. 

ASMFC 2007. Special Report to the ASMFC Atlantic Sturgeon Management Board: Estimation 

of Atlantic sturgeon bycatch in the coastal Atlantic commercial fisheries of New England 

and the mid-Atlantic. Washington (DC): Atlantic States Marine Fisheries Commission. 

Aadland LP, Cook C. 1992. An electric trawl for sampling bottom dwelling fishes in deep turbid 

streams. North America Journal of Fisheries Management 12:62-656. 

Agusa T, Kunito T, Tanabe S, Pourkazemi M, Aubrey DG. 2004. Concentrations of trace 

elements in muscle of sturgeon in the Caspian Sea. Marine Pollution Bulletin 49:789-800. 

Alam SK, Brim MS, Carmody GA, Parauka FM. 2000. Concentrations of heavy and trace metals 

in muscle and blood of juvenile Gulf sturgeon (Acipenser oxyrinchus desotoi) from the 

Suwannee River, Florida. Journal of Environmental Science and Health A35:645-660. 

American Fisheries Society (AFS). 2004. Guidelines for the use of fishes in research: American 

Fisheries Society, American Institute of Fishery Research Biologists, American Society 

of Ichthyologists and Herpetologists. Bethesda (MD): American Fisheries Society.

Website address: http://www.fisheries.org/afs/docs/policy_16.pdf

Apperson KA, Anders PJ. 1991. Kootenai River white sturgeon investigations and experimental 

culture. Annual Progress Report FY 1990 to Bonneville Power Administration. Portland 

(OR): Idaho Department of Fish and Game. 

Atlantic Sturgeon Status Review Team (ASSRT). 2007. Status Review of Atlantic sturgeon 

(Acipenser oxyrinchus oxyrinchus). Gloucester (MA): Report to National Marine 

Fisheries Service, Northeast Regional Office. Web address: 

http://www.nmfs.noaa.gov/pr/pdfs/statusreviews/atlanticsturgeon2007.pdf 

Bain MB. 1997. Atlantic and shortnose sturgeon of the Hudson River: common and divergent 

life history attributes. Environmental Biology of Fishes 48:347-358. 

Baker DW, Peake SJ, Kieffer JD. 2008. The effect of capture, handling, and tagging on 

hematological variables in wild adult lake sturgeon. North American Journal of Fisheries 

Management 28:296-300. 

Barton BA, Dwyer WP. 1997. Physiological stress effects of continuous- and pulsed-DC on 

electroshock on juvenile bull trout. Journal of Fish Biology 51:998-1008. 

Bath DW, O’Connor JM, Alber JB, Arivdson LG. 1981. Development and identification of 

larval Atlantic sturgeon (Acipenser oxyrinchus) and shortnose sturgeon (A. brevirostrum) 

from the Hudson River estuary, New York. Copeia 1981:711-717. 
50

Bauer C, Unfer G, Loupal G. 2005. Potential problems with external trailing antennae: antenna 

migration and ingrowth of epithelial tissue, a case study from a recaptured Chondrostoma 

nasus. Journal of Fish Biology 67:885-889. 

Berg J. 2006. A review of contaminant impacts on the Gulf of Mexico sturgeon, Acipenser 

oxyrinchus desotoi. Report to United States Fish and Wildlife Service, Panama City, 

Florida. 

Billard R, Cosson J, Noveiri SB, Porkazemi M. 2004. Cryopreservation and short-term storage of 

sturgeon sperm, a review. Aquaculture 236:1-9. 

Birstein V J, Bemis WE, Waldman JR. 1997. The threatened status of acipenseriform species: a 

summary. Environmental Biology of Fishes 48:427-435. 

Bordner CE, Doroshov SI, Hinton DE, Pipkin RE, Fridley RB, Haw F. 1990. Evaluation of 

marking techniques for juvenile and adult white sturgeon reared in captivity. American 

Fisheries Society 7:293-303. 

Borisenko AV, Lim BK, Ivanova NV, Hanner RH, Hebert PDN. 2008. DNA barcoding in 

surveys of small mammal communities: a field study in Suriname. Molecular Ecology 

Resources 8(3):471-479. 

Brennan JS, Cailliet GM. 1989. Comparative age-determination techniques for white sturgeon in 

California. Transactions of the American Fisheries Society 118:296-310. 

Brosse L, Dumont P, Lepage M, Rochard E. 2002. Evaluation of a gastric lavage 

method for sturgeon. North American Journal of Fisheries Management 22:955-960. 

Bruch RM, Dick TA, Choudhury A. 2001. A field guide for the identification of stages of gonad 

development in lake sturgeon, (Acipenser fulvescens Rafinesque), with notes on lake 

sturgeon reproductive biology and management implications. Oshkosh (WI): Wisconsin 

Department of Natural Resources. 

Brundage HM. 2003. Report to the National Marine Fisheries Service, Northeast Regional 

Office: Contaminant analysis of tissues from a shortnose sturgeon (Acipenser 

brevirostrum) from the Kennebec River, Maine. Chadds Ford (PA): Environmental 

Research and Consulting, Inc. 

Bryan JL, Wildhaber ML, Papoulias DM, DeLonay AJ, Tillitt DE, Annis ML. 2007. Estimation 

of gonad volume, fecundity, and reproductive stage of shovelnose sturgeon using 

sonography and endoscopy with application to the endangered pallid sturgeon. Journal of 

Applied Ichthyology 23:411-419. 

Buckley J, Kynard B. 1985. Yearly movements of shortnose sturgeon in the Connecticut River. 

Transactions of the American Fisheries Society 114:813-820. 
51

Buddington RK, Christofferson JP. 1985. Digestive and feeding characteristics of the 

chondrosteans. Environmental Biology of Fishes 14:31-41. 

Carr SH, Tatman F, Chapman FA. 1996. Observations on the natural history of the Gulf of 

Mexico sturgeon (Acipenser oxyrinchus desotoi Vladykov 1955). Ecology of Freshwater 

Fish 5:169-174. 

Chapman FA, Van Eenennaam JP, Doroshov SI. 1996. The reproductive condition of white 

sturgeon (Acipenser transmontanus), in San Francisco Bay, California. Fishery Bulletin 

94:628-634. 

Chapman FA, Park C. 2005. Comparison of sutures used for wound closure in sturgeon 

following a gonad biopsy. North American Journal of Aquaculture 67:98-101. 

Chebanov MS, Galich EV. 2009. Ultrasound diagnostics in sturgeon broodstock management. 

6-th International symposium on sturgeon 2009, Wuhan, China: Workshop on sturgeon 

sexing and gonad staging, 47 p. 

Collins MR, Smith TIJ. 1993. Characteristics of the adult segment of the Savannah River 

population of shortnose sturgeon. Proceedings of the Annual Conference of the 

Southeastern Association of Fish and Wildlife Agencies 47:485-491. 

Collins MR, Smith TIJ, Heyward LD. 1994. Effectiveness of six methods for marking juvenile 

shortnose sturgeon. Progressive Fish-Culturist 56:250-254. 

Collins MR, Smith TIJ. 1996. Sturgeon fin ray removal is nondeleterious. North American 

Journal of Fisheries Management 16:939-941. 

Collins MR, Rogers SG, Smith TIJ. 1996. Bycatch of sturgeons along the southern Atlantic coast 

of the USA. North American Journal of Fisheries Management 16:24-29. 

Colombo RE, Wills PS, Garvey JE. 2004. Use of ultrasound imaging to determine sex of 

shovelnose sturgeon. North American Journal of Fisheries Management 24:322-326. 

Conte FS, Doroshov SI, Lutes PB, Strange EM. 1988. Hatchery manual for the white sturgeon 

(Acipenser transmontanus Richardson) with application to other North American 

Acipenseridae. Oakland (CA): University of California Cooperative Extension Service 

Publication 3322. 

Counihan TD, Miller AI, Mesa MG, Parsley MJ. 1998. The effects of dissolved gas 

supersaturation on white sturgeon larvae. Transactions of the American Fisheries Society 

127:316-322. 

Crocker CE, Cech, JJ Jr. 1996. The effects of hypercapnia on the growth of juvenile white 

sturgeon, Acipenser transmontanus. Aquaculture 147:293–299. 
52

Cuerrier JP. 1951. The use of pectoral fin rays for determining age of sturgeon and other species 

of fish. Canadian Fish Culturist 11:10-18. 

Dadswell MJ, Taubert BD, Squiers TS, Marchette D, Buckley J. 1984. Synopsis of biological 

data on shortnose sturgeon, Acipenser brevirostrum LeSueur 1818. Washington (DC): 

NOAA Technical Report NMFS 14. 

Dick TA, Jarvis SR, Sawatzky CD, Stewart DB. 2006. The lake sturgeon, Acipenser fulvescens

(Chondrostei: Acipenseridae): An annotated bibliography. Winnipeg (Manitoba): 

Canadian Technical Report of Fisheries and Aquatic Sciences 2671. 

Dickson W. 1989. Cod gill net simulation model. Fisheries Research 7:149-174. 

DiLauro MN, Krise WF, Hendrix MA, Baker SE. 1994. Short-term storage of Atlantic sturgeon 

sperm. The Progressive Fish-Culturist 56:143-144. 

Dorsey KM. 2009. Quality assessment of Atlantic Sturgeon (Acipenser oxyrinchus) spermatozoa 

under conditions of short-term storage [master’s thesis]. [College Park (MD)]: University 

of Maryland. 

Dovel WL, Berggren TJ. 1983. Atlantic sturgeon of the Hudson Estuary, New York. New York 

Fish and Game Journal 30:140-172 

Dovel WL, Pekovitch AW, Berggren TJ. 1992. Biology of the shortnose sturgeon (Acipenser 

brevirostrum Lesueur, 1818) in the Hudson River estuary, New York. In Smith CL, 

editor. Estuarine Research in the 1980s. Albany (NY): State University of New York 

Press, p. 187-216. 

Doyle W, Paukert C, Starostka A, Hill T. 2008. A comparison of four types of sampling gear 

used to collect shovelnose sturgeon in the Lower Missouri River. Journal of Applied 

Ichthyology 24:637-642. 

Edwards RE, Parauka FM, Sulak KJ. 2007. New insights into marine migration and winter 

habitat of Gulf sturgeon. In: Anadromous sturgeons: Habitats, threats, and management. 

Munro J, Hatin D, Hightower JE, McKown K, Sulak KJ, Kahnle AW, Caron F, editors. 

Bethesda (MD): American Fisheries Society. p. 183-196. 

Elliott J, Beamesderfer R. 1990. Comparison of efficiency and selectivity of gears used to 

sample white sturgeon in a Columbia River reservoir. California Fish and Game 76:174-

180. 

Erickson DL, Hightower JE. 2007. Oceanic distribution and behavior of green sturgeon. 

American Fisheries Society Symposium 56:197-211. 
53

Erickson DL, Kahnle A, Millard MJ, Mora EA, Bryija G, Higgs A, Mohler J, DuFour M, 

Kenney G, Sweka J, Pikitch EK. [In Review]. Use of pop-up satellite archival tags to 

identify oceanic-migratory patterns for adult Atlantic sturgeon, Acipenser oxyrinchus 

oxyrinchus. Journal of Applied Ichthyology. 

Eyler S, Mangold M, Minkkinen S. 2009. Atlantic coast sturgeon tagging database. Annapolis 

(MD): US Fish and Wildlife Service. Summary Report, February 2009. 

Fiest G, Van Eenennaam JP, Doroshov SI, Schreck CB, Schneider RP, Fitzpatrick MS. 2004. 

Early identification of sex in cultured white sturgeon, Acipenser transmontanus, using 

plasma steroid levels. Aquaculture 232:581-590. 

Foster JR. 1977. Pulsed gastric lavage: An efficient method of removing the stomach contents of 

live fish. Progressive Fish-Culturist 39:166-169. 

Fox DA, Hightower JE, Parauka FM. 2000. Gulf sturgeon spawning migration and habitat in the 

Choctawhatchee River system, Alabama-Florida. Transactions of the American Fisheries 

Society 129:811-826. 

Fuller SA, Henne J, Seals J, Mudrak V. 2008. Performance of commercially available Passive 

Integrated Transponder (PIT) tag systems used for fish identification and 

interjurisdictional fisheries management. North American Journal of Fisheries 

Management 28:386-393. 

Geoghegan P, Mattson MT, Keppel RG. 1992. Distribution of the shortnose sturgeon in the 

Hudson River estuary, 1984-1988. In Smith CL, editor. Estuarine research in the 1980s. 

Albany (NY): State University of New York Press. p. 217-277. 

Golder Associates Ltd. 2006a. Columbia River adult white sturgeon capture, transport and 

handling manual, May 2006 Version. Castlegar (British Columbia): Golder Associates 

Ltd. Prepared for Upper Columbia White Sturgeon Recovery Initiative. 

Golder Associates Ltd. 2006b. Upper Columbia River juvenile white sturgeon monitoring: 

Phase 4 investigations, 2005 – 2006. Castlegar (British Columbia): Golder Associates, 

LTD. Report prepared for BC Hydro, Castlegar, B.C. Golder Report No. 05-1480-058F. 

Gray CA, Johnson DD, Broadhurst MK, Young DJ. 2005. Seasonal, spatial, and gear-related 

influences on relationships between retained and discarded catches in multi-species gill 

net fishery. Fisheries Research 75:56-72. 

Hager C. 2007. Sink gill net fisheries and description of factors that can contribute to higher or 

lower interaction and retention rates in estimation of Atlantic sturgeon bycatch in coastal 

Atlantic commercial fisheries of New England and the mid-Atlantic, A special report to 

ASMFC Atlantic Sturgeon Technical Committee. 
54

Haley NJ. 1998. A gatric lavage technique for characterizing diets of sturgeon. North American 

Journal of Fisheries Management 18:978-981. 

Hamley JM. 1975. Review of gill net selectivity. J. Fish. Res. Board Can. 32:1943-1969. 

Hanner R, Corthals A, Dessauer HC. 2005. Salvage of genetically valuable tissues following a 

freezer failure. Molecular Phylogenetics and Evolution 34:425-455. 

Harms CA, Bakal RS. 1994. Techniques in fish anesthesia. American Association of Zoo 

Veterinarians Annual Proceedings 1994:202–210. 

Hartley WG. 1967. Electronarcosis of fish for handling. In Vibert R, editor. Fishing with 

electricity: its application to biology and management. London (UK): Fishing News 

Books. P. 251-255. 

Helm WT, Tyus HM. 1992. Influence of coating type on retention of dummy transmitters 

implanted in rainbow trout. North America Journal of Fisheries Management 12:257–

259. 

Henyey E., Kynard B, Zhuang P. 2002. Use of electronarcosis to immobilize juvenile lake and 

shortnose sturgeons for handling and the effects on their behavior. Journal of Applied 

Ichthyology 18:502-504. 

Hilton EJ, Bemis WE. 1999. Skeletal variation in shortnose sturgeon (Acipenser brevirostrum) 

from the Connecticut River: implications for comparative osteological studies of fossil 

and living fishes. In Arratia G, Schultze HP, editors. Mesozoic Fishes 2 – Systematics 

and Fossil Record. Verlag Dr. Friedrich Pfeil, München(Germany): Verlag Dr. Friedrich 

Pfeil. P. 69-94 

Hilton EJ. 2002. Observations on the rostral canal bones of two species of Acipenser

(Actinopterygii, Acipenseriformes). Copeia 2002:213–219. 

Hoff TB, Klauda RJ, Young JR. 1988. Contributions to the biology of shortnose sturgeon in the 

Hudson River Estuary. In Smith CL, editor. Fisheries research in the Hudson River. 

Albany (NY): State University of New York Press. P. 171-189. 

Holst R, Wileman D, Madsen N. 2002. The effect of twine thickness on the size selectivity and 

fishing power of Baltic cod gill nets. Fisheries Research 56:303–312. 

Holliman FM, Reynolds JB. 2002. Electroshock-induced injury in juvenile white sturgeon. North 

American Journal of Fisheries Management 22:494-499. 

Horvath A, Wayman WR, Urbanyi B, Ware KM, Dean JC, Tiersch TR. 2005. The relationship of 

the cryoprotectants methanol and dimethyl sulfoxide and hyperosmotic extenders on 

sperm cryopreservation of two North-American sturgeon species. Aquaculture 247:243-

251. 
55

Hovgard H, Lassen H. 2000. Manual on estimation of selectivity for gill net and longline gears in 

abundance surveys. Rome (Italy): FAO Fisheries Technology, Paper. 

Hrabik RA, Herzog DP, Ostendorf DE, Petersen MD. 2007. Larvae provide first evidence of 

successful reproduction by pallid sturgeon, Scaphirhynchus albus, in the Mississippi 

River. Journal of Applied Ichthyology 23:436-443. 

Huff JA. 1975. Life history of Gulf of Mexico sturgeon, Acipenser oxyrinchus desotoi, in 

Suwannee River, Florida. St. Petersburg (FL): Florida Department of Natural Resources, 

Marine Research Laboratory No. 16. 

Hurty CA, Diaz DE, Campbell JL, Lewbart GA. 2001. Chemical analysis of six commercial 

adult iguana, Iguana iguana, diets. Journal of Herpetological Medicine and Surgery 

11(3):23-26. 

Isely JJ, Fontenot QC. 2000. Retention of coded wire tags in juvenile shortnose sturgeon. North 

American Journal of Fisheries Management 20:1040-1043. 

Jähnichen H, Warnecke D, Trölsch E, Kohlmann K, Bergler H, Pluta HJ. 1999. Motility and 

fertilizing capability of cryopreserved Acipenser ruthenus L. sperm. Journal of Applied 

Ichthyology 15:204-206. 

Jenkins WE, Smith T, Heyward L, Knott M. 1993. Tolerance of shortnose sturgeon, Acipenser 

brevirostrum, juveniles to different salinity and dissolved oxygen concentrations. 

Proceedings of the Southeast Association of Fish and Wildlife Agencies 47:476–484. 

Kieffer M, Kynard B. 1993. Annual movements of shortnose and Atlantic sturgeons in the 

Merrimack River, Massachusetts. Transactions of the American Fisheries Society 

122:1088-1103. 

Knights BC, Lasee BA. 1996. Effects of implanted transmitters on adult bluegill at two 

temperatures. Transactions of the American Fisheries Society 125:440—449. 

Kobayashi T, Takeuchi Y, Takeuchi T, Yoshizaki G. 2007. Generation of viable fish from 

cryopreserved primordial germ cells. Molecular Reproduction and Development 74:207-

213. 

Kohlhorst DW. 1976. Sturgeon spawning in the Sacramento River in 1973 as determined by 

distribution of larvae. California Fish and Game 62:32-40. 

Kruse GO, Scarnecchia DL. 2002. Assessment of bioaccumulated metal and organochlorine 

compounds in relation to physiological biomarkers in Kootenai River white sturgeon. 

Journal of Applied Ichthyology 18:430-438. 
56

Kynard B, Kieffer M, Burlingame M, Hogan M. 1999. Studies on shortnose sturgeon. Amherst 

(MA): University of Massachusetts, Final Report to Northeast Utilities Service Co., 

Berlin, CT. 

Kynard B, Kieffer M. 2002. Use of a borescope to determine the sex and egg maturity stage of 

sturgeons and the effect of borescope use on reproductive structures. Journal of Applied 

Ichthyology 18:505-508. 

Light RW, Adler PH, Arnold DE. 1983. Evaluation of gastric lavage for stomach analyses. North 

American Journal of Fisheries Management 3:81-85. 

Livia L, Antonella P, Hovirag L, Mauro N, Panara F. 2006. A nondestructive, rapid, reliable and 

inexpensive method to sample, store and extract high-quality DNA from fish body mucus 

and buccal cells. Molecular Ecology Notes 6:257-260. 

MacDonald DD, Ikonomou MG, Rantalaine AL, Rogers IH, Southerland D, Van Oostdam J. 

1997. Contaminants in white sturgeon (Acipenser transmontanus) from the Upper Fraser 

River, British Columbia, Canada. Environmental Toxicology and Chemistry 16:479-499. 

Machiels MAM, Klinge M, Lanters R, van Densen WLT. 1994. Effect of snood length and 

hanging ratio efficiency and selectivity of bottom-set gill nets for pikeperch Stizostedion 

lucioperca L. and bream Abramis brama. Fisheries Research 9:231-239. 

Marais JFK. 1985. Some factors influencing the size of fishes caught in gill nets in eastern cape 

estuaries. Fisheries Research 3:251-261. 

Marchant SR, Shutters MK. 1996. Artificial substrates collect Gulf sturgeon eggs. North 

American Journal of Fisheries Management 16:445-447. 

Matsche M, Bakal R. 2008. General and reproductive health assessments of shortnose sturgeon 

with application to Atlantic sturgeon: anesthesia, phlebotomy, and laparoscopy. Oxford 

(MD): Maryland Department of Natural Resources. 

Mbogori MN, Kimani M, Kuria A, Lagat M, Danson JW. 2006. Optimization of FTA technology 

for large scale plant DNA isolation for use in marker assisted selection. African Journal 

of Biotechnology 5:693-696. 

McCabe GT, Beckman LG. 1990. Use of an artificial substrate to collect white sturgeon eggs. 

California Fish and Game 76:248-250. 

McCleave JD, Fried SM, Towt AK. 1977. Daily movements of shortnose sturgeon, Acipenser 

brevirostrum, in a marine estuary. Copeia 1977:149-157. 

McCord JW. 1998. Investigation of fisheries parameters for anadromous fisheries in South 

Carolina. Charleston (SC): South Carolina Department of Natural Resources. Completion 

report to National Marine Fisheries Service (AFC-53). 
57

Meehan WR, Miller RA. 1978. Stomach flushing: effectiveness and influence on survival and 

condition of juvenile salmonids. Journal of the Fisheries Research Board of Canada 

35:1359-1363. 

Miller TJ. 2007. Factors associated with mortality of incidentally caught sturgeon in the 

Northwest Atlantic Ocean in estimation of Atlantic sturgeon bycatch in coastal Atlantic 

commercial fisheries of New England and the mid-Atlantic. A Special Report to ASMFC 

Atlantic Sturgeon Technical Committee. 

Moghim M, Vajhi AR, Veshkini A, Masoudifard M. 2002. Determination of sex and maturity in 

Acipenser stellatus by using ultrasonography. Journal of Applied Ichthyology 18:325-

328. 

Mohler JW. 1994. Evaluation of tagging procedures and associated tag retention in hatcheryproduced yearling of Atlantic sturgeon (Acipenser oxyrhinchus). Lamar (PA): USFWS 

Northeast Fishery Center Fiscal Year 1994 Annual Report. Study number L-94-03. 

Mohler JW. 2004. Culture manual for the Atlantic sturgeon, Acipenser oxyrinchus oxyrinchus. 

Hadley (MA): US Fish and Wildlife Service Region 5. Web address: 

http://www.fws.gov/northeast/fisherycenter/pdfs/ASTCultureManual.pdf

Moser ML, Ross SW. 1993. Distribution and movements of shortnose sturgeon (Acipenser 

brevirostrum) and other anadromous fishes of the lower Cape Fear River, North Carolina. 

Wilmington (NC): Final Report to the US Army Corps of Engineers. 

Moser ML, Ross SW. 1995. Habitat use and movements of shortnose and Atlantic sturgeons in 

the lower Cape Fear River, North Carolina. Transactions of the American Fisheries 

Society 124:225-234. 

Moser ML, Bichy JB, Roberts SB. 1998. Sturgeon distribution in North Carolina. Wilmington 

(NC): Final report to the US Army Corps of Engineers, Wilmington District. 

Moser ML, Bain M, Collins MR, Haley N, Kynard B, O’Herron II JC, Rogers G, Squiers TS. 

2000. A protocol for use of shortnose and Atlantic sturgeon. Silver Spring (MD): NOAA 

Technical Memorandum NOAA Fisheries Service-OPR-18. 

Murphy MD, Heagey RF, Neugebauer VH, Gordon MD, Hintz JL. 1995. Mortality of spotted 

seatrout released from gill-net or hook-and-line gear in Florida. North American Journal 

of Fisheries Management 15:748-753. 

Musick JA, Jenkins RE, Burkhead NB. 1994. Sturgeons, family Acipenseridae. In: Jenkins RE, 

Burkhead NB, editors. Freshwater Fishes of Virginia. Bethesda (MD): American 

Fisheries Society. P. 183-190. 
58

Neiffer DL, Stamper MA. 2009. Fish sedation, anesthesia, analgesia, and euthanasia: 

Considerations, methods, and types of drugs. ILAR Journal 50(4):343-360. 

Niklitschek EJ. 2001. Bioenergetics modeling and assessment of suitable habitat for juvenile 

Atlantic and shortnose sturgeons (Acipenser oxyrinchus and A. brevirostrum) in the 

Chesapeake Bay [dissertation]. [College Park(MD)]: University of Maryland. 

Niklitschek EJ, Secor DH. 2005. Modeling spatial and temporal variation of suitable nursery 

habitats for Atlantic sturgeon in the Chesapeake Bay. Estuarine, Coastal and Shelf 

Science 64:135–148. 

Niklitschek EJ, Secor DH. 2009. Dissolved oxygen, temperature, and effects on the 

ecophysiology and survival of juvenile Atlantic sturgeon in estuarine waters: I. 

Laboratory results. Journal of Experimental Marine Biology and Ecology 

381(Supplement 1):S150-S160. 

O’Herron JC, Able KW. 1990. A study of the endangered shortnose sturgeon (Acipenser 

brevirostrum) in the Delaware River. New Brunswick (NJ): NJ Department of 

Environmental Protection. Final Performance Report to US Fish and Wildlife Service and 

New Jersey Fish, Game, and Wildlife, Department of Environmental Protection. Project 

AFS-10-R. 

Paragramian VL, Beamesderfer RCP. 2003. Growth estimates from tagged white sturgeon 

suggest that ages from fin rays underestimate true ages in the Kootenai River, USA and 

Canada. Transactions of the American Fisheries Society 132:895-903. 

Park C, Chapman FA. 2005. An extender solution for the short-term storage of sturgeon semen. 

North American Journal of Aquaculture 67:52-57. 

Peterson DL, Bain MB, Haley N. 2000. Evidence of declining recruitment of Atlantic sturgeon in 

the Hudson River. North American Journal of Fisheries Management 20:231-238. 

Pine WE, Flowers HJ, Johnson KG, Jones ML. 2006. An assessment of Gulf sturgeon 

movement, spawning site selection, and post-spawn holding areas in the Apalachicola 

River, Florida. Gainesville (FL): University of Florida. Final report to FWRI Freshwater 

Fisheries Research, Requisition no. PR640342, 104 p. 

 

Purbayanto A, Tsunoda A, Akiyama S, Arimoto T, Tokai T. 2000. Survival of Japanese whiting 

Sillago japonica and by-catch species captured by a sweeping trammel net. Fisheries 

Science 67:21–29. 

Purvis LB, Villegas P, Perozo F. 2006. Evaluation of FTA® paper and phenol for storage, 

extraction and molecular characterization of infectious bursal disease virus. Journal of 

Virological Methods 138:66-69. 
59

Rien TA, Beamesderfer RC. 1994. Accuracy and precision of white sturgeon age estimates from 

pectoral fin rays. Transactions of the American Fisheries Society 123:255-265. 

Rien TA, Beamesderfer RCP, Foster CA. 1994. Retention, recognition, and effects on survival of 

several tags and marks for white sturgeon. California fish and Game 80(4):161-170. 

Reis EG, Pawson MG. 1999. Fish morphology and estimating selectivity by gill nets. Fisheries 

Research 39:263-273. 

Reynolds JB. 1996. Electrofishing. In Murphy BR, Willis DW, editors. Fisheries Techniques, 

2nd edition. Bethesda (MD): American Fisheries Society. p. 221-253. 

RL & L Environmental Services Ltd. 1996. Columbia River white sturgeon investigations. 1995 

study results. Vancouver (BC): BC Ministry of Environment, Lands and Parks, Nelson 

Region. Report Prepared for B.C. Hydro, Kootenay Generation RL & L Report No. 96-

377F. 

Rossiter A, Noakes DL, Beamish FWH. 1995. Validation of age estimation for the lake sturgeon. 

Transactions of the American Fisheries Society 124:777-781. 

Savoy T. 1991. Sturgeon status in Connecticut waters. Waterford (CT): Connecticut Department 

of Environmental Protection, Completion Report AFC-18-3. 

Savoy T. 2007. Prey eaten by Atlantic sturgeon in Connecticut waters. American Fisheries 

Society Symposium 56:157-165. 

Savoy T, Benway J. 2004. Food habits of shortnose sturgeon collected in the lower Connecticut 

River from 2000 through 2002. In Jacobson PM, Dixon DA, Leggett WC, Marcy BC 

2nd., Massengill RR, editors. The Connecticut River Ecological Study (1965-1973) 

revisited: ecology of the lower Connecticut River 1973-2003. Bethesda (MD): American 

Fisheries Society, Monograph 9. p. 353-360. 

Scott WB, Crossman EJ. 1973. Freshwater fishes of Canada. Fisheries Research Board of 

Canada, Bulletin 184:1-966. 

Seaburg KG. 1957. A stomach sampler for live fish. Progressive Fish-Culturist 19:137-139. 

Secor DH, Niklitschek EJ. 2001. Hypoxia and sturgeons: Report to the Chesapeake Bay Program 

dissolved oxygen criteria team. Solomons (MD): University of MD Center for 

Environmental Science. Technical Report Series No. TS-314-01-CBL. 

Secor DH, Woodland R. 2005. Recovery and status of shortnose sturgeon in the Hudson River. 

Solomons (MD): University of MD Center for Environmental Science. Report to Hudson 

River Foundation for Science and Environmental Research, Inc. HRF 011/02A, UMCESCBL 07-4-35255. 
60

Shuman DA, Peters EJ. 2007. Evaluation of pulsed gastric lavage on the survival of captive 

shovelnose sturgeon. Journal of Applied Ichthyology 23:521-524. 

Smith TIJ, Lamprecht SD, Hall JW. 1990. Evaluation of tagging techniques for shortnose 

sturgeon, Acipenser brevirostrum, and Atlantic sturgeon A. oxyrhynchus. American 

Fisheries Society Symposium 7:134-141. 

Snyder DE. 1988. Description and identification of shortnose and Atlantic sturgeon larvae. 

American Fisheries Society Symposium 5:7-30. 

Sprague CR, Beckman LG, Duke SD. 1993. Prey selection by juvenile whitesturgeon in 

reservoirs of the Columbia River. In: Beamesderfer RC, Nigro AA, editors. Status and 

habitat requirements of the white sturgeon populations in the Columbia River 

downstream from McNary Dam, vol. 2. Portland (OR): Oregon Department of Fish and 

Wildlife. Final Report to Bonneville Power Administration. p. 229-243. 

Stein AB, Friedland KD, Sutherland M. 2004. Atlantic sturgeon marine bycatch mortality on the 

continental shelf of the northeast United States. North American Journal of Fisheries 

Management 24(1):171-183. 

Stevenson JT, Secor DH. 1999. Age determination and growth of Hudson River Atlantic 

sturgeon, Acipenser oxyrinchus. Fishery Bulletin 97:153-166. 

Stoskopf MK. 1993. Clinical physiology. In: Stoskopf MK, editor. Fish medicine. Philadelphia 

(PA): W.B. Saunders Company. p. 48-57. 

Sulak KJ, Clugston JP. 1998. Early life history stages of gulf sturgeon in the Suwannee River, 

Florida. Transactions of the American Fisheries Society 127:758-771. 

Sweka JA, Mohler JW, Millard MJ. 2006. Relative abundance sampling of juvenile Atlantic 

sturgeon in the Hudson River. Lamar (PA): US Fish and Wildlife Service. Final Study 

Report to the New York Department of Environmental Conservation, Hudson River 

Fisheries Unit.

Taubert BD. 1980. Reproduction of shortnose sturgeon (Acipenser brevirostrum) in Holyoke 

Pool, Connecticut River, Massachusetts. Copeia 1980:114-117. 

Van Eenennaam JP, Doroshov SI. 1998. Effects of age and body size on gonadal development in 

Atlantic sturgeon (Acipenser oxyrinchus mitchill). Journal of Fish Biology 53:624-637. 

Vecsei P, Litvak MK, Noakes DG, Rien T, Hochleithner M. 2003. A noninvasive technique for 

determining sex of live adult North American sturgeons. Environmental Biology of 

Fishes 68:333-338. 

Vladykov VD, Greeley JR. 1963. Order Acipenseroidei. In: Bigelow HB, Breder CM, Cohen 

DM, Mead GW, Merriman D, Olsen YH, Schroeder WC, Schultz LP, Tee-Van J, editors. 
61

Fishes of the Western North Atlantic, memoir 1, part three, of the Sears Foundation for 

Marine Research. New Haven (CT): Yale University. 

Walker MK, Yanke EA, Gingerich WH. 1994. Use of electronarcosis to immobilize juvenile and 

adult northern pike. The Progressive Fish-Culturist 56:237-243. 

Wanner GA. 2006. Evaluation of a gastric lavage method on juvenile pallid sturgeon. North 

American Journal of Fisheries Management 26:587-591. 

Webb MAH, Feist GW, Foster EP, Schreck CB, Fitzpatrick MS. 2002. Potential classification of 

sex and stage of gonadal maturity of wild white sturgeon using blood plasma indicators. 

Transactions of the American Fisheries Society 131:132-142. 

Webb MAH, Erickson DL. 2007. Reproductive structure of the adult green sturgeon, Acipenser 

medirostris, population in the Rogue River, Oregon. Environmental Biology of Fishes 

79:305-314. 

Wei Q, Kynard B. 1996. Study of Chinese sturgeon spawning in the Yangtze River. Jingshao 

(Hubei Province): Yangtze River Fishery Research Institute. Progress Report. 

Whiteman KW, Travnichek VH, Wildhaber ML, DeLonay A, Papoulias D, Tillett D. 2004. Age 

estimates for shovelnose sturgeon: a cautionary note based on annulus formation in 

pectoral fin rays. North American Journal of Fisheries Management 24:731-734. 

Wildhaber ML, Papoulias DM, Delonay AJ, Tillitt DE, Bryan JL, Annis ML, Allert JA. 2005. 

Gender identification of shovelnose sturgeon using ultrasonic and endoscopic imagery 

and the application of the method to the pallid sturgeon. Journal of Fish Biology 67:114-

132. 

Wildhaber ML, Papoulias DM, Delonay AJ, Tillitt DE, Bryan JL, Annis ML. 2006. 

Development of methods to determine the reproductive status of pallid sturgeon in the 

Missouri River. Columbia (MO): US Geological Survey. Final Report to the US Fish and 

Wildlife Service. 

Wilkie MP, Davidson K, Brobbel MA, Kieffer JD, Booth RK, Bielak AT, Tufts BL. 1996. 

Physiology and survival of wild Atlantic salmon following angling in warm summer 

waters. Transactions of the American Fisheries Society125:572-580. 

Wingate RL, Secor D. 2007. Intercept telemetry of the Hudson River striped bass resident 

contingent: migration and homing patterns. Transactions of the American Fisheries 

Society 136:95-104. 

Yokota K, Fujimori Y, Shiode D, Tokai T. 2001. Effect of thin twine on gill net size-selectivity 

analyzed with the direct estimation method. Fisheries Science 67:851-856. 
62

Zehfuss KP, Hightower JP, Pollock KH. 1999. Transactions of the American Fisheries Society 

128:130-143. 

Zehfuss KP. 2000. The status, movement, habitat preference and monitoring of Gulf sturgeon in 

several Florida rivers [dissertation]. [Raleigh (NC)]: North Carolina State University. 

Ziegeweid JR. 2006. Ontogenetic changes in salinity and temperature tolerances of young-ofthe-year shortnose sturgeon, Acipenser brevirostrum [thesis]. [Athens (GA)]: University 

of Georgia. 

Ziegeweid JR, Jennings CA, Peterson DL. 2008a. Thermal maxima of juvenile shortnose 

sturgeon acclimated to different temperatures. Environmental Biology of Fishes 82:299-

307. 

Ziegeweid JR, Jennings CA, Peterson DL, Black MC. 2008b. Effects of salinity, temperature, 

and weight on the survival of young-of-year shortnose sturgeon. Environmental Biology 

of Fishes 137:1490-1499. 
63

APPENDIX – FIELD DATA SHEET 
64
Publishing in NOAA Technical Memorandum NMFS-NE

Manuscript Qualification

This series represents a secondary level of scientific

publishing in the National Marine Fisheries Service (NMFS).

For all issues, the series employs thorough internal scientific

review, but not necessarily external scientific review. For most

issues, the series employs rigorous technical and copy editing.

Manuscripts that may warrant a primary level of scientific

publishing should be initially submitted to one of NMFS's

primary series (i.e., Fishery Bulletin, NOAA Professional Paper

NMFS, or Marine Fisheries Review).

Identical, or fundamentally identical, manuscripts should

not be concurrently submitted to this and any other publication

series. Manuscripts which have been rejected by any primary

series strictly because of geographic or temporal limitations

may be submitted to this series.

Manuscripts by Northeast Fisheries Science Center

(NEFSC) authors will be published in this series upon approval

by the NEFSC's Deputy Science & Research Director. Manuscripts by non-NEFSC authors may be published in this series

if: 1) the manuscript serves the NEFSC's mission; 2) the

manuscript meets the Deputy Science & Research Director's

approval; and 3) the author arranges for the printing and binding

funds to be transferred to the NEFSC's Research Communications Branch account from another federal account. For all

manuscripts submitted by non-NEFSC authors and published

in this series, the NEFSC will disavow all responsibility for the

manuscripts' contents; authors must accept such responsibility.

The ethics of scientific research and scientific publishing

are a serious matter. All manuscripts submitted to this series

are expected to adhere -- at a minimum -- to the ethical guidelines

contained in Chapter 2 ("Publication Policies and Practices") of

the Scientific Style and Format: the CSE Manual for Authors,

Editors, and Publishers, seventh edition (Reston VA: Council

of Science Editors). Copies of the manual are available at

virtually all scientific libraries.

edition of the United States Government Printing Office Style

Manual. That style manual is silent on many aspects of scientific

manuscripts. NEFSC publication and report series rely more on the

CSE Style Manual, seventh edition.

For in-text citations, use the name-date system. A special

effort should be made to ensure that the list of cited works contains

all necessary bibliographic information. For abbreviating serial

titles in such lists, use the guidance of the International Standards

Organization; such guidance is easily accessed through the various

Cambridge Scientific Abstracts’ serials source lists (see http://

www.public.iastate.edu/~CYBERSTACKS/JAS.htm). Personal communications must include date of contact and full name and mailing

address of source.

For spelling of scientific and common names of fishes, mollusks, and decapod crustaceans from the United States and Canada,

use Special Publications No. 29 (fishes), 26 (mollusks), and 17

(decapod crustaceans) of the American Fisheries Society (Bethesda

MD). For spelling of scientific and common names of marine

mammals, use Special Publication No. 4 of the Society for Marine

Mammalogy (Lawrence KS). For spelling in general, use the most

recent edition of Webster’s Third New International Dictionary of

the English Language Unabridged (Springfield MA: G. & C.

Merriam).

Typing text, tables, and figure captions: Text, tables, and

figure captions should be converted to Word. In general, keep text

simple (e.g., do not switch fonts and type sizes, do not use hard

returns within paragraphs, do not indent except to begin paragraphs). Also, do not use an automatic footnoting function; all notes

should be indicated in the text by simple numerical superscripts, and

listed together in an "Endnotes" section prior to the "References

Cited" section. Especially, do not use a graphics function for

embedding tables and figures in text.

Tables should be prepared with a table formatting function.

Each figure should be supplied in digital format (preferably GIF or

JPG), unless there is no digital file of a given figure. Except under

extraordinary circumstances, color will not be used in illustrations.

Manuscript Preparation

Organization: Manuscripts must have an abstract, table

of contents, and -- if applicable -- lists of tables, figures, and

acronyms. As much as possible, use traditional scientific

manuscript organization for sections: "Introduction," "Study

Area," "Methods & Materials," "Results," "Discussion" and/

or "Conclusions," "Acknowledgments," and "References Cited."

Style: All NEFSC publication and report series are

obligated to conform to the style contained in the most recent

Northeast Fisheries Science Center

Operations, Management & Information Division

Research Communications Branch

Editorial Office

Manuscript Submission

Authors must submit separate digital files of the manuscript

text, tables, and figures. The manuscript must have cleared

NEFSC's online internal review system. Non-NEFSC authors who

are not federal employees will be required to sign a "Release of

Copyright" form.

Send all materials and address all correspondence to: Jarita A.

Davis (Editor), Editorial Office, NMFS Northeast Fisheries Science Center, 166 Water Street, Woods Hole, MA 02543-1026.
National Marine Fisheries Service, NOAA

166 Water St.

Woods Hole, MA 02543-1026

Publications and Reports

of the

Northeast Fisheries Science Center

The mission of NOAA's National Marine Fisheries Service (NMFS) is "stewardship of living marine resources 

for the benefit of the nation through their science-based conservation and management and promotion of the 

health of their environment." As the research arm of the NMFS's Northeast Region, the Northeast Fisheries 

Science Center (NEFSC) supports the NMFS mission by "conducting ecosystem-based research and assessments of living marine resources, with a focus on the Northeast Shelf, to promote the recovery and long-term 

sustainability of these resources and to generate social and economic opportunities and benefits from their use." 

Results of NEFSC research are largely reported in primary scientific media (e.g., anonymously-peer-reviewed 

scientific journals). However, to assist itself in providing data, information, and advice to its constituents, the 

NEFSC occasionally releases its results in its own media. Currently, there are three such media:

NOAA Technical Memorandum NMFS-NE -- This series is issued irregularly. The series typically includes: data reports of 

long-term field or lab studies of important species or habitats; synthesis reports for important species or habitats; annual reports 

of overall assessment or monitoring programs; manuals describing program-wide surveying or experimental techniques; literature 

surveys of important species or habitat topics; proceedings and collected papers of scientific meetings; and indexed and/or annotated 

bibliographies. All issues receive internal scientific review and most issues receive technical and copy editing.

Northeast Fisheries Science Center Reference Document -- This series is issued irregularly. The series typically includes: data 

reports on field and lab studies; progress reports on experiments, monitoring, and assessments; background papers for, collected

abstracts of, and/or summary reports of scientific meetings; and simple bibliographies. Issues receive internal scientific review, but 

no technical or copy editing.

Resource Survey Report (formerly Fishermen's Report) -- This information report is a quick-turnaround report on the distribution 

and relative abundance of selected living marine resources as derived from each of the NEFSC's periodic research vessel surveys

of the Northeast's continental shelf. There is no scientific review, nor any technical or copy editing, of this report.

OBTAINING A COPY: To obtain a copy of a NOAA Technical Memorandum NMFS-NE or a Northeast Fisheries Science Center 

Reference Document, or to subscribe to the Resource Survey Report, either contact the NEFSC Editorial Office (166 Water St., 

Woods Hole, MA 02543-1026; 508-495-2228) or consult the NEFSC webpage on "Reports and Publications" (http://www.nefsc.

noaa.gov/nefsc/publications/).

ANY USE OF TRADE OR BRAND NAMES IN ANY NEFSC PUBLICATION OR REPORT DOES NOT IMPLY ENDORSEMENT.

MEDIA

 MAIL