A Protocol for 

Use of Shortnose, Atlantic, 

Gulf, and Green Sturgeons 

Jason Kahn and Malcolm Mohead 

U.S. Department of Commerce 

National Oceanic and Atmospheric Administration 

National Marine Fisheries Service 

NOAA Technical Memorandum NMFS-OPR-45 

March 2010 
Atlantic sturgeon (Robert Michelson, Photography by Michelson, Inc.) 

Gulf sturgeon (Oscar Sosa, New York Times) 

Green sturgeon (Thomas Dunklin) 

Cover: shortnose sturgeon (credit: Robert Michelson)
A Protocol for 

Use of Shortnose, Atlantic, 

Gulf, and Green Sturgeons 

Jason Kahn and Malcolm Mohead 

NOAA Technical Memorandum NMFS-OPR-45 

March 2010

U.S. Department of Commerce 

Gary Locke, Secretary of Commerce 

National Oceanic and Atmospheric Administration 

Jane Lubchenco, Ph.D., Administrator of NOAA 

National Marine Fisheries Service 

Eric C. Schwaab, Assistant Administrator for Fisheries
ii 

Suggested citation: 

Kahn, Jason, and Malcolm Mohead. 2010. A Protocol for Use of Shortnose, Atlantic, 

Gulf, and Green Sturgeons. U.S. Dep. Commerce, NOAA Tech. Memo. NMFS-OPR-45, 

62 p. 

A copy of this report may be obtained from: 

Jason Kahn 

Office of Protected Resources 

NMFS, NOAA 

1315 East West Highway 

Silver Spring, MD 20910 

jason.kahn@noaa.gov 
iii 

TABLE OF CONTENTS 

INTRODUCTION ........................................................................................................................................ 1

NON-TARGETED SPECIES CONCERNS IN THE RESEARCH AREA ............................................. 2

CAPTURE ..................................................................................................................................................... 3

DISSOLVED OXYGEN, TEMPERATURE, AND SALINITY ............................................................................ 3

GILLNETS AND TRAMMEL NETS ............................................................................................................... 6

ELECTROFISHING ...................................................................................................................................... 8

OTHER NON-LETHAL SAMPLING GEAR................................................................................................... 8

TRAWLING................................................................................................................................................. 9

D-NETS ...................................................................................................................................................... 9

EGG MATS ............................................................................................................................................... 10

OTHER METHODS OF EGG COLLECTION ............................................................................................... 10

RECOMMENDATIONS .............................................................................................................................. 10

HANDLING AND HOLDING .................................................................................................................. 12

PROPER HANDLING OF STURGEON ........................................................................................................ 12

SHORT-TERM HOLDING ......................................................................................................................... 12

RECOMMENDATIONS .............................................................................................................................. 14

STANDARD RESEARCH METHODS ................................................................................................... 15

MEASURING ............................................................................................................................................. 15

WEIGHING ............................................................................................................................................... 16

PHOTOGRAPHING .................................................................................................................................... 16

PIT TAGS ................................................................................................................................................ 16

GENETIC TISSUE SAMPLING ................................................................................................................... 18

RECOMMENDATIONS .............................................................................................................................. 18

ANESTHETIZATION ............................................................................................................................... 20

CHEMICAL ANESTHETIC ........................................................................................................................ 22

PHYSICAL ANESTHETIC .......................................................................................................................... 25

RECOMMENDATIONS .............................................................................................................................. 27

TAGGING ................................................................................................................................................... 28

TELEMETRY TAGS .................................................................................................................................. 28

EXTERNAL IDENTIFIER TAGS ................................................................................................................. 31

RECOMMENDATIONS .............................................................................................................................. 32

GASTRIC LAVAGE .................................................................................................................................. 33

RECOMMENDATIONS .............................................................................................................................. 35

SEX IDENTIFICATION ........................................................................................................................... 36

ENDOSCOPY ............................................................................................................................................. 36

SURGICAL BIOPSY ................................................................................................................................... 38

ULTRASOUND .......................................................................................................................................... 39

BLOOD PLASMA ...................................................................................................................................... 40

RECOMMENDATIONS .............................................................................................................................. 41

AGE ESTIMATION .................................................................................................................................. 42

ACCURACY AND PRECISION OF ESTIMATES .......................................................................................... 43

AGE VALIDATION .................................................................................................................................... 45

DELETERIOUS EFFECTS OF FIN SPINE SAMPLING ................................................................................. 46

ALTERNATIVE METHODS FOR AGE ESTIMATION .................................................................................. 46
iv 

RECOMMENDATIONS .............................................................................................................................. 47

SALVAGE SPECIMENS .......................................................................................................................... 48

ACKNOWLEDGEMENTS ....................................................................................................................... 49

REFERENCES ........................................................................................................................................... 50
1 

Introduction 

The goal of the National Marine Fisheries Service (NMFS) protocols for the use of 

sturgeon is standardization of research practices to benefit the recovery of Gulf of 

Mexico (Gulf), green, Atlantic, and shortnose sturgeon while also minimizing potentially 

negative impacts of research. As with A Protocol for the Use of Shortnose and Atlantic 

Sturgeon (Moser et al. 2000a), these protocols provide guidelines for consistent and safe 

sampling methods when conducting research on sturgeon. They were developed from a 

comprehensive review of the best available scientific information at the time of 

publication, including peer reviewed journals, technical memorandums, species status 

reviews, interviews with researchers, and empirical evidence provided by researchers. 

Currently, some state agencies have been delegated authority for issuing research permits 

for Gulf and green sturgeon. However, due to previous lack of protocols established for 

these species, they were incorporated into this document. 

The majority of research conducted on sturgeon falls into several categories: capturing, 

handling, holding, standard research, anesthetization, tagging, gastric lavage, sex 

identification and stage of maturation, and age estimation. First, sturgeon must be 

captured, which may also require consideration of the waterway sampled to mitigate 

impacts on other federally listed threatened or endangered species. NMFS has 

determined that measuring, passive integrated transponder (PIT) tagging, and genetic 

sampling are essential procedures to provide NMFS with the most basic information on 

each fish and therefore those procedures are strongly recommended. After those 

procedures are completed, other discretionary research might include telemetry tagging, 

gastric lavage, sex identification, and age estimation. These discretionary procedures 

should use either chemical or physical anesthesia, potentially increasing risks to sturgeon. 

These protocols were developed to allow for safe, non-lethal research on sturgeon, 

balancing the necessary negative impacts of research while still allowing researchers to 

gather information vital to the recovery of listed species under the Endangered Species 

Act (ESA). These protocols are based on a thorough and comprehensive review of the 

best available scientific information on current research methods and the subsequent risk 

to these species. When researchers or managers have reason to exceed recommendations 

in this document using less known or riskier techniques, NMFS recommends first using 

surrogate Acipenserids or hatchery-reared sturgeon. When researchers or managers feel 

non-recommended methods must be conducted on wild listed or candidate species, the 

researchers should consult with the appropriate permitting agency in order to justify why 

their methodology is necessary to provide information for the recovery of these species. 
2 

Non-Targeted Species Concerns in the Research Area 

When sampling shortnose, Atlantic, Gulf, and green sturgeon, the potential exists for 

researchers to encounter other ESA or Marine Mammal Protection Act (MMPA) listed 

species, in addition to other locally or state protected species. These circumstances will 

vary with location and NMFS encourages consultation with the appropriate management 

authority in all cases. 

When other ESA protected species are potentially present in an action area, the researcher 

must contact NMFS or the US Fish and Wildlife Service (USFWS) for clarification on 

the likelihood to adversely impact any listed species, or destroy or adversely modify any 

critical habitat for that species. The presence of listed species may require researchers to 

alter sampling plans to avoid taking listed fish, such as Pacific or Atlantic salmonids, or 

mammals, such as Stellar sea lions or manatees. 

In many other locations, marine mammals, protected under the MMPA but not the ESA, 

may be present. The MMPA places a moratorium, with certain exceptions, on the taking 

and importing of marine mammals and marine mammal products. In 1981, Congress 

amended the MMPA to allow the incidental, but not intentional, taking of small numbers 

of marine mammals by U.S. citizens who engage in a specified activity (other than 

commercial fishing) within a specified geographical region. If marine mammals, 

including non-ESA listed pinnipeds or cetaceans, have the potential to be taken incidental 

to scientific research activities on sturgeon (e.g., there is a chance of entanglement), the 

researcher should consult with NMFS under section 101(a)(5) of the MMPA to determine 

if an incidental take authorization is warranted. Contact: Office of Protected Resources, 

Silver Spring, Maryland (301-713-2289). 

In other instances, predators may frequent sampling areas posing threats to listed 

sturgeon species. In such cases, nets must be monitored at all times and pulled if 

predators are evidenced. Pinnipeds have been seen feeding on listed sturgeon by 

researchers (Fernandez 2008, Marty Gingras, California Department of Fish and Game, 

pers. comm.), potentially other predatory species such as odontocetes and sharks could 

take sturgeon while trapped in gillnets or trammel nets. If there are reasons to believe 

sturgeon could be harmed by predators while captured in gillnets or trammel nets, those 

nets should be continuously monitored. 
3 

Capture 

Researchers most often capture Gulf, Atlantic, green, and shortnose sturgeon using a 

variety of gears including gillnets (drift and anchored), trammel nets, seine nets, trawls, 

trot lines, pound nets, and electrofishing. Nets of varying length and mesh size are 

chosen to target different life stages of sturgeon (Mason and Clugston 1993, DeVries 

2006). 

Generally, sturgeon are hardy, allowing some research methods lethal to other fish. 

These methods can still be stressful to sturgeon, occasionally resulting in lethal and, more 

often, sub-lethal effects. For example, during pre-spawning activities, capture and 

handling is thought to have resulted in immediate downstream migration or aborted 

spawning runs (Moser and Ross 1995, Kynard et al. 2007, Gail Wippelhauser, Maine 

Department of Marine Resources, pers. comm.). Also, during periods of warm water or 

low dissolved oxygen (DO), fish have been lethally stressed (Hastings et al. 1987, Secor 

and Gunderson 1998). NMFS recommends capturing adult sturgeon while they are still 

in their winter staging areas, but does not recommend targeting sturgeon during their 

upstream spawning migration due to the risks of aborted spawning runs. However, when 

the purpose of the research is to document the size of the spawning run, managers must 

determine whether the information to be gained is worth the risk posed by the research. 

Dissolved Oxygen, Temperature, and Salinity 

For all sturgeon species, research has revealed that survival is affected by a relationship 

between temperature, DO, and salinity and this vulnerability may be increased by the 

research-related stress of capture, holding, and handling. The following environmental 

information is considered relevant for establishing recommendations for directed 

sampling on early life stages to adult life stages of sturgeon. 

Jenkins et al. (1993), Secor and Gunderson (1998), Niklitschek (2001), Secor and 

Niklitschek (2001 and 2002), and Niklitshek and Secor (2009a and 2009b) demonstrated 

shortnose and Atlantic sturgeon survival in a laboratory setting was affected by reduced 

DO, increased temperature, or increased salinity. Other researchers have demonstrated 

similar relationships between temperature, DO, and salinity in green sturgeon (Van 

Eenennaam et al. 2005, Allen et al. 2006, Allen and Cech 2007). Likewise, Altinok et al.

(1998), Sulak and Clugston (1998), Sulak and Clugston (1999), and Waldman et al.

(2002) reported high temperatures, low DO, and high salinities result in lower survival of 

Gulf sturgeon. 

Though there may be differences between populations in different geographical regions, 

optimal growth for both Atlantic and shortnose sturgeon has been shown to occur at 70% 

oxygen saturation with a temperature of approximately 20°C (Niklitschek 2001). 

Shortnose sturgeon have also been shown to experience significant reductions in food 

consumption when temperatures exceed 25.8°C (Niklitschek 2001). Green sturgeon 

require cooler temperatures, growing optimally between 15° and 19°C, and experiencing 

reduced growth rates between 20° and 24°C (Mayfield and Cech 2004). However, larval 

green sturgeon grow more optimally at 24°C compared to 19°C (Allen et al. 2006). Gulf 
4 

sturgeon also appear dependent on temperature for optimal growth, fasting during hot 

summer months and feasting during winter when water temperatures and DO in the Gulf 

of Mexico and tributaries are more optimal (Sulak and Randall 2002). 

Considerable work has been conducted on temperature tolerances of sturgeon (Wang et 

al. 1985, Wehrly 1995, Kynard 1997, Campbell and Goodman 2004, Cech and Doroshov 

2004, Van Eenennaam et al. 2005, Ziegeweid et al. 2007, Sardella et al. 2008). In recent 

work on critical thermal maximum, Ziegeweid et al. (2007) demonstrated hatchery-raised 

young of year shortnose sturgeon can tolerate between 28° and 30°C, while the maximum 

safe temperature limits for adults ranges between 28° and 31°C. Kynard (1997) also 

notes empirical temperatures of 28° to 30°C in summer months creates unsuitable 

shortnose sturgeon habitat. Atlantic sturgeon experience lower survival when water 

temperatures exceed 28°C (Niklitshek and Secor 2005). Mayfield and Cech (2004) 

estimated the lethal water temperature for green sturgeon in the wild at 27°C. Sardella et 

al. (2008) found green sturgeon lethal limits in a laboratory is approximately 33°C, in 

freshwater and sea water, although the maximum respiratory response evidenced is 26° to 

28°C. Although Gulf sturgeon reside in freshwater during summer months where water 

temperatures range from 28° to 32°C, there have been no studies estimating lethal 

temperature limits for Gulf sturgeon. It is worth noting, however, the healthiest 

population of Gulf sturgeon occurs in the Suwannee River, where temperatures are 

generally maintained at 28°C by springs in parts of the river. 

There is no clear evidence to suggest minimum water temperatures negatively affect 

sturgeon when captured beyond the early life stages. Therefore, this document identifies 

only upper water temperature restrictions to establish safe sampling limits for threatened 

or endangered sturgeon. However, when air temperatures are below freezing, handling 

procedures should be limited to less than two minutes to prevent exposure of a sturgeon’s 

skin to freezing temperatures. 

Because warm water can hold less DO, percent oxygen saturation is a measurement that 

accounts for water temperatures and DO concentrations, providing a general index of 

how much DO is available to sturgeon under various environmental conditions. All three 

measures are used in this document to highlight risks to sturgeon survival (Table 1). The 

24 hour LC50 (concentration lethal to 50% of the test fish) of DO for shortnose sturgeon 

is documented between 2.2 and 3.1 mg/L at temperatures ranging from 22°C to 29°C 

(Campbell and Goodman 2004). Secor and Niklitschek (2002) reported the critical DO 

concentration for Eurasian sturgeons to be 4.5 mg/L at 24°C, but also found 3.6 mg/L DO 

critical at 20°C. Following a similar pattern, critical concentrations of DO between 4.3 

and 4.7 mg/L were found for shortnose and Atlantic sturgeon at temperatures ranging 

from 22° and 27°C respectively. Further, acute lethal effects to shortnose and Atlantic 

sturgeon were observed when DO was 3.3 mg/L at temperatures between 22° and 27°C 

(Secor and Niklitschek 2002). Survival of Atlantic sturgeon was observed to be 100% in 

water temperatures of 26°C with 7 mg/L DO; however, 12% survival was observed in 

waters with 3 mg/L DO at the same temperature (Secor and Gunderson 1998). Even 

when water temperatures were only 19°C and DO was 3 mg/L, 25% of the Atlantic 

sturgeon died. Similar to reduced growth rates experienced by shortnose sturgeon when 
5 

temperatures are above 25°C, both shortnose and Atlantic sturgeon growth is impaired 

when DO is less than 4.7 mg/L (Secor and Niklitschek 2002). Jenkins et al. (1993) 

confirmed 12% mortality for 339 mm juvenile sturgeon when held at 2.5 mg/L DO and 

22.5°C, while no sturgeon died when DO was above 4 mg/L at any temperature. 

Likewise, Secor and Gunderson (1998) found the DO level required avoiding mortality 

was 5 mg/L. Specific DO tolerance levels have not been established for green or Gulf 

sturgeon, although hypoxia for many Acipenser species has been documented to begin at 

4 mg/L (Cech et al. 1984, Jenkins et al. 1993, Secor and Gunderson 1998). Similarly, 

Cech and Crocker (2002) identified hypoxia for sturgeon as 58% oxygen saturation. 

Table 1. Water temperature, dissolved oxygen, percent oxygen saturation of the water, 

and survival rates of sturgeon tested. 

Authors Species Temp (°C) DO (mg/L) % Saturation Effects 

Jenkins et al.

1993 Shortnose 22.5 2.5 29% 88% survival 

Campbell and 

Goodman 

2004 

Shortnose 22 – 29 2.2 – 3.1 25 - 41% 50% survival 

Secor and 

Niklitschek 

2002 

Atlantic 

and 

shortnose 

22 – 27 3.3 38 – 42% Acute lethal 

effects 

Secor and 

Gunderson 

1998 

Atlantic 26 3 37% 12% survival 

Secor and 

Gunderson 

1998 

Atlantic 19 3 33% 75% survival 

Secor and 

Niklitshek 

2002 

Eurasian 

24 4.5 54% Critical DO 

concentration, 

onset of sublethal effects 20 3.6 40% 

Secor and 

Niklitshek 

2002 

Atlantic 

and 

shortnose 

22 – 27 4.3 – 4.7 50 – 60% 

Critical DO 

concentration, 

onset of sublethal effects 

NMFS recognizes the synergistic effects of water temperature and DO present difficulties 

when establishing finite levels for safe sturgeon sampling (Table 1). It is clear from 

reported empirical catch data and scientific literature, higher temperatures and lower DOs 

stress sturgeon even if the percent oxygen saturation remains constant or increases. 

Water temperature and DO can be responsible for mortality events. Each individual 

sturgeon will react differently to changes in environmental conditions such as water 

quality, salinity, and stress associated with capture and handling, which compounds the 

difficulty of conducting a risk assessment. 
6 

Using data reported from capture of shortnose and Atlantic sturgeon from the 1970s to 

present and the critical thresholds and LC50s reported in the scientific literature as 

reference points, NMFS established safe environmental limits for capturing and handling 

sturgeon species. NMFS recommends not capturing or handling Gulf, Atlantic and 

shortnose sturgeon when DO concentrations are below 4.5 mg/L. Green sturgeon should 

not be captured or handled when DO concentrations are below 5 mg/L. Additionally, 

NMFS recommends not sampling for Gulf, shortnose, or Atlantic sturgeon when 

temperatures exceed 28°C and green sturgeon should not be captured when water 

temperatures exceed 25°C. When establishing these recommendations, NMFS also 

considered the percent oxygen saturation of water and recommends not sampling for 

Gulf, Atlantic, or shortnose sturgeon when oxygen saturation is below 55% or green 

sturgeon when oxygen saturation is below 58%. Sampling at higher temperatures or 

lower DO levels may be possible if the percent oxygen saturation in water is maintained 

at these levels. 

Gillnets and Trammel Nets 

Researchers typically use gillnets and trammel nets to capture sturgeon. These netting 

techniques, while potentially lethal for many species of fish, are somewhat safer for 

sturgeon. However, given the implications of water temperature, DO, and percent 

oxygen saturation, both soak times and mesh size are important factors considered for 

safely capturing and handling sturgeon. Mesh size that is too small for the targeted life 

stage is more likely to constrict gills resulting in mortality via suffocation. The mesh size 

chosen for gill netting sturgeon, therefore, should be carefully considered and appropriate 

for the species and life stage targeted. Experimental nets with multiple mesh sizes may 

be appropriate for researchers to discover the safest and most effective mesh size. For 

example, due to disproportionately high reports of mortality using ten inch stretch mesh 

with Atlantic sturgeon (Balazik et al. 2009), this size mesh should not be used to sample 

adult Atlantic or Gulf sturgeon. 

Safe net soak times are influenced by water temperature, DO, and, to a lesser extent, 

salinity. While there are no publications documenting the effects of soak times on 

mortality rates of sturgeon, there is consensus amongst sturgeon researchers that shorter 

soak times are safer than longer soak times (Mark Collins, South Carolina Department of 

Natural Resources; Matt Fisher, Delaware Division of Fish and Wildlife; Dewayne Fox, 

Delaware State University; Chris Hager, Virginia Institute of Marine Science; Doug 

Peterson, University of Georgia; William Post, South Carolina Department of Natural 

Resources; Mike Randall, United States Geological Survey (USGS); and Ken Sulak, 

USGS, pers. comm.). By monitoring signs of stress such as excessive redness, mucous 

production, or lethargy, experienced researchers will often shorten net deployment 

regardless of measured environmental conditions (Kathryn Hattala, New York State 

Department of Environmental Conservation; Tom Savoy, Connecticut Department of 

Environmental Protection; and Doug Peterson, University of Georgia, pers. comm.). 

When using anchored gillnets while targeting Atlantic and shortnose sturgeon, soak times 

of 14 hours are safe when water temperatures at the sampling depth are under 15°C. 

However, soak times should not exceed four hours in waters up to 20°C, two hours in 
7 

waters up to 25°C, and one hour in waters up to 28°C at the sampling depth (Table 2). 

Similar effects were alluded to in Moser et al. (2000a), but were not clearly defined. 

Gulf sturgeon net set durations should not exceed four hours under any conditions. 

Mortalities have been documented in the empirical records of researchers while fishing 

above 20°C at net set durations ranging from 45 minutes to 24 hours. However, 

mortalities have been extremely rare when fishing nets less than two hours and at 

temperatures between 20° and 25°C. The one hour soak time at water temperatures 

between 25° and 28°C (Table 2) accommodates standard research practices of netting at 

slack tides (i.e., the occurrence of relatively still water at the turn of the low tide). There 

have been only two recorded sturgeon mortalities documented when fishing in this 

manner. 

Table 2. Appropriate fishing protocols for Gulf, Atlantic, and shortnose sturgeon. 

Net set duration 

(hours) 

Temperature at 

sampling depth 

Minimum DO at 

sampling depth 

% oxygen saturation 

at sampling depth 

14† Up to 15°C 4.5 mg/L 55% 

4 15° to 20°C 4.5 mg/L 55% 

2 20° to 25°C 4.5 mg/L 55% 

1 25° to 28°C 4.5 mg/L 55% 

No sampling Over 28°C 4.5 mg/L 55% 

† Net set duration for Gulf sturgeon should not exceed four hours for all temperatures up to 20°C. 

When fishing for green sturgeon, NMFS recommends that gill net fishing not be 

conducted in the Sacramento River, California all year to prevent interactions with listed 

salmonids and to also protect green sturgeon during their upstream migrations. NMFS 

also recommends that no gillnetting or trammel netting occur in the Feather River 

between October 31st and March 1st of each year to protect spawning salmonids. When 

fishing for green sturgeon in other locations, the risk of interactions between gillnets or 

trammel nets and listed salmonids or pinnipeds requires the nets to be manned at all 

times. Additionally, pinnipeds are protected by the MMPA and the presence of gillnets 

in the water could pose an entanglement risk and require an Incidental Take 

Authorization (Section 101(a)(5) of the MMPA). NMFS recommends net soak times 

should not exceed four hours in water temperature up to 19°C, two hours between 19° 

and 23°C, and one hour for water temperature between 23° and 25°C (Table 3). 

Table 3. Appropriate fishing protocols for green sturgeon. 

Net soak times 

(hours) 

Temperature at 

sampling depth 

Minimum DO at 

sampling depth 

% oxygen saturation 

at sampling depth 

4 Up to 19°C 5 mg/l 58% 

2 19° to 23°C 5 mg/l 58% 

1 23° to 25°C 5 mg/l 58% 

No netting Over 25°C 5 mg/l 58% 

When following the protocols in Table 2 between 2005 and 2009, East Coast sturgeon 

researchers recorded over 3,800 captures of shortnose sturgeon resulting in no mortality. 
8 

However, while fishing outside of these recommended criteria, the same researchers 

experienced a 0.6% mortality rate of captured shortnose sturgeon. This is the same 

mortality rate documented for shortnose sturgeon captured between 2000 and 2004 when 

researchers followed the Moser et al. (2000a) protocols. 

When drift gillnetting, nets are allowed to drift on the rising tide or in slack tide until just 

after high tide for approximately thirty minutes to several hours, depending on the 

location and swiftness of the tide. Water quality conditions and net soak times for drift 

gill nets are the same as for anchored gillnets. However, drift nets must be tended 

because of the risk of gear entanglement or loss of gear resulting in ghost nets. For drift 

gillnet fishing, gear should be pulled immediately if it is obvious a sturgeon has been 

captured. 

Electrofishing 

Electrofishing gear poses documented risks and potentially lethal effects to all sturgeon 

species (Moser et al. 2000b, Holliman and Reynolds 2002). Sturgeon have exceptional 

electro-sensory abilities and actively avoid electrofishing gear (Moser et al. 2000b). If 

sturgeon are likely present in areas where agencies are using electrofishing gear to target 

other species, only low voltage direct current should be used if no alternative sampling 

method is available. While electrofishing likely reduces feeding and alters spawning 

behavior (Moser et al. 2000b), such sub-lethal effects may not be significantly different 

than effects caused by other capture methods. However, due to more effective and safer 

methods of capture, NMFS prohibits electrofishing to capture Gulf, green, Atlantic, or 

shortnose sturgeon. 

Other Non-Lethal Sampling Gear 

While fyke, hoop, and pound nets are not commonly used by researchers to capture 

sturgeon, they occasionally capture sturgeon as bycatch in several fisheries. Usually 

sturgeon captured as bycatch in these gear types are found in relatively good condition. 

Large numbers of sturgeon captured in fyke, hoop, and pound nets have been used by 

researchers in cooperation with these commercial fisheries in Canada. Because these nets 

are less stressful to sturgeon, they are an acceptable alternative to gillnets. 

Set lines have also been used to effectively sample white, pallid, shovelnose, and lake 

sturgeon and are approved options for sampling Gulf, green and Atlantic sturgeon as 

well. Shortnose sturgeon are less likely to be taken on a set line because of their diets. 

The two concerns with set lines are predation and hooking mortality. If there are 

predators such as pinnipeds in the area, the set line should be monitored constantly and 

pulled if any predators are seen surfacing. The hooks can be swallowed, damaging 

organs such as the gills and stomach, if the hook sizes are too large or small for the 

targeted sturgeon life stage. Every effort should be made to limit and monitor adverse 

effects, including not using set lines in some locations if they cannot be fished without 

mortality. 
9 

Trawling 

While gillnets and trammel nets are most commonly used for targeting adult and subadult sturgeon, they are not as effective as trawls at capturing young of the year juvenile 

sturgeon. In larger river systems such as the Mississippi and Missouri River, and more 

recently in Atlantic coastal rivers, researchers have successfully employed a modified 

“Missouri trawl” (Herzog et al. 2005) — a two-seam (i.e., standard) slingshot balloon 

trawl (Gutreuter et al. 1995) completely covered with heavy, delta-style mesh. 

Trawls in general are limited by shallow water (less than 20 inches) and benthic 

obstacles. The location of trawling should be monitored using a sounding device and 

global positioning system to avoid snags and limit repeated disturbance of the same 

location. The tow rope should be quickly released from the boat if any debris is caught 

and the trawl unengaged to minimize damage to the substrate or catch. Ideally, a chase 

boat is recommended to assist with recovery of the cod end or assisting with snags, but if 

that is not possible, a buoy should be attached to a single 70 to 100 foot rope line fastened 

to the cod end of the trawl to assist retrieval if the trawl becomes snagged. 

The footrope of a trawl should maintain contact with the substrate during conditions of 

heavy current, fast tow speeds, or undulating bottom surfaces (e.g., sand waves). The 

trawl should be operated attached to the boat with 100 to 200 foot towlines, the length 

dependent on water depth (i.e., deeper water required longer towlines as reported in 

Brabant and Nedelec 1979). The trawl should be manually deployed and retrieved by 

powering the boat in reverse (bow upstream) with continued movement downstream. A 

standard haul should be approximately 300 to 500 feet, lasting approximately 10 minutes, 

and towed at a range of three to five knots (Gutreuter et al. 1995). 

Areas successful for trawling are characterized by a variety of habitat substrate including 

fine and course sands with mobile bedforms (sand dunes) and mudflats. Particularly 

productive areas are located at the mouths of tributaries entering a larger river. However, 

any large, straight river segment, devoid of benthic material that may entangle nets, can 

be successfully trawled. 

D-Nets 

When targeting eggs and early life stage (ELS) sturgeon, the two commonly used 

sampling methods are D-nets and artificial substrates. Both techniques can be non-lethal, 

but due to the risk of mortality, no more eggs and ELS sturgeon should be captured than 

are absolutely necessary. While not mandatory, in rivers with unknown spawning 

populations, adults can be tagged and tracked to document possible spawning runs and 

spawning areas prior to sampling for eggs (Kieffer and Kynard 1996). Otherwise, D-nets 

should be deployed well before the earliest time spawning would be expected. Due to the 

risks associated with capturing and impinging ELS sturgeon in the D-Nets, however, they 

should be checked at least every three hours to minimize incidental mortality (Boyd 

Kynard, USGS, pers. comm.). D-nets should also be equipped with flow meters to 

calculate filtered water volume when developing an index of abundance and spawning 

success (# ELS/ volume of water sampled) (Taubert 1980). If the purpose of the research 

is to verify the occurrence of spawning, nets should be checked every hour. As soon as 
10 

ELS are captured, sampling should be discontinued. If the purpose of the research is to 

verify duration of the spawning period, then additional samples may need to be taken, but 

the acceptable number of ELS fish to be captured would depend on the status of the 

sturgeon populations in the river. 

Egg Mats 

Artificial substrates consist of floor buffing pads or similar materials, approximately two 

feet in diameter (described in Fox et al. 2000) for the purpose of collecting eggs as they 

are deposited in the water column. These pads should be anchored to the river bottom in 

suspected spawning areas. No more pads should be fished than is necessary. If the 

researcher is unsure of the number of pads required to identify spawning areas and 

success, no more than 100 to 150 pads should be fished at once across several sites. Pads 

should be checked at least twice a week or more frequently if circumstances allow. The 

artificial substrates should be examined in the field for sturgeon eggs and only returned to 

the river if more samples are needed. If it is not necessary to remove the eggs from the 

mat, the mat can be returned to the river bottom allowing the eggs to incubate and hatch 

before being removed. For every artificial substrate that collects an egg, environmental 

conditions such as latitude, longitude, velocity, substrate type, depth, dissolved oxygen, 

etc. should be collected.

Other Methods of Egg Collection 

There are other methods of sampling eggs and ELS, such as epibethic sleds, 

ichthyoplankton nets, and pump sampling. These methods are not considered as effective 

as the other described methods, though they are acceptable sampling methods. 

Recommendations 

General 

• NMFS recommends capturing adult sturgeon while they are still in their winter 

staging areas, but does not recommend targeting sturgeon during their upstream 

spawning migration due to the risks of aborted spawning runs. 

Water Temperature, Dissolved Oxygen, and Salinity 

• When air temperatures are below freezing, handling procedures should be limited 

to less than two minutes to prevent exposure of a sturgeon’s skin to freezing 

temperatures. 

• NMFS recommends Gulf, Atlantic, and shortnose sturgeon are not captured or 

handled when DO concentrations are below 4.5 mg/L. Green sturgeon should not 

be captured or handled when DO concentrations are below 5 mg/L. 

• NMFS recommends not sampling for Gulf, shortnose, or Atlantic sturgeon occur 

when temperatures exceed 28°C; while sampling for green sturgeon should not 

occur when temperatures exceed 25°C. 

• NMFS recommends not sampling for Gulf, Atlantic, or shortnose sturgeon when 

the oxygen saturation is below 55% and not sampling green sturgeon when the 

oxygen saturation is below 58%. 
11 

Gillnets and Trammel Nets 

• Due to disproportionately high reports of mortality using ten inch stretch mesh 

with Atlantic sturgeon, this size mesh should not be used to sample adult Atlantic 

or Gulf sturgeon. 

• NMFS recommends no gill net fishing be conducted in the Sacramento River, 

California all year round to prevent interactions with listed salmonids and also to 

protect green sturgeon during their upstream migrations. 

• NMFS also recommends that no gillnetting or trammel netting take place in the 

Feather River, California between October 31st and March 1st of each year to 

protect spawning salmonids. 

• NMFS recommends net soak times should not exceed four hours in water 

temperature up to 19°C, should not exceed two hours between 19° and 23°C, and 

one hour for water temperature between 23° and 25°C. 

• Gillnets should be used sparingly and carefully in waters where other listed 

species may be encountered. The researcher must contact NMFS or the USFWS 

when other listed species may be incidentally affected. 

Electrofishing 

• NMFS prohibits electrofishing to capture Gulf, green, Atlantic, or shortnose 

sturgeon. 

Other Non-Lethal Sampling Gear 

• Fyke, hoop, and pound nets are an acceptable alternative to gillnets for Gulf, 

green, Atlantic, and shortnose sturgeon. 

• Set lines are approved options for sampling Gulf, green, and Atlantic sturgeon. 

Trawling 

• NMFS recommends trawling as safe, efficient sampling gear to target small 

juvenile Gulf, Atlantic, shortnose, and green sturgeon; however, small mesh 

gillnets and trammel nets are also acceptable. 

D-Nets 

• NMFS recommends D-nets and egg mats to sample rivers for eggs or ELS of 

Gulf, Atlantic, shortnose, or green sturgeon. 

• Due to risks associated with capturing and impinging ELS sturgeon in D-Nets, 

they should be checked at least every three hours to minimize incidental 

mortality. 

Egg Mats 

• No more egg mats should be fished than is necessary. If the researcher is unsure 

of the number of pads required to identify spawning areas and success, no more 

than 100 to 150 pads should be fished at once. 
12 

Handling and Holding 

Handling of sturgeon refers to the time period actual research activities are conducted on 

live fish and does not refer to the time a fish is held in live cars before and after research 

activities. Holding is the period of time a sturgeon is in possession but kept in live cars 

either waiting to be handled or recovered from handling prior to being released. 

Proper Handling of Sturgeon 

Improper handling can result in lethal or sub-lethal impacts to sturgeon. In some cases, 

sturgeon may display altered behavior after being released, for example, swimming 

towards the ocean rather than remaining in the river, or, in some instances, aborting 

spawning runs completely (Moser and Ross 1995, Schaffter 1997, Kelly et al. 2007, 

Benson et al. 2007, Moser and Lindley 2007). There are no other alternatives to handling 

sturgeon during research; however, the researcher’s primary focus should be the wellbeing of the sturgeon. 

NMFS strongly recommends standard handling procedures performed on all sturgeon 

captured including measuring, weighing, PIT tagging, and tissue sampling. The total 

time required to complete routine research procedures should not exceed 15 minutes. 

Additional procedures such as internal tagging, lavage, boroscoping, etc. will take more 

time for handling and recovery. However, only one additional discretionary procedure to 

the standard handling procedures should be performed on each sturgeon, thus minimizing 

handling time prior to release. For example, if a sturgeon is fitted with a telemetry tag, it 

should not also undergo gastric lavage. And when water temperatures are above 23°C for 

green sturgeon or 25°C for Gulf, shortnose, or Atlantic sturgeon, the extent of research 

should be limited to the standard handling procedures of measuring, weighing, PIT 

tagging, and tissue sampling. 

Fish should be handled rapidly, but with care and kept in water to the maximum extent 

possible during handling. During handling procedures, each fish should be immersed in a 

continuous stream of ambient water passing over the sturgeon’s gills. Many sturgeon 

researchers provide sturgeon with supplemental compressed oxygen, thereby reducing 

stress and ensuring DO does not fall below acceptable saturation levels. 

Researchers should also attempt to support larger sturgeon in slings preventing struggle 

during transfer. Sturgeon should be weighed using hand held sling scales or a platform 

scale for larger sturgeon. Also, because sturgeon are sensitive to direct sunlight, they 

should be covered and kept moist. 

Short-Term Holding 

All captured sturgeon should be removed from the capture gear and immediately 

transferred to short-term holding. When multiple fish are captured, those not processed 

immediately should be held in a net pen or live car while waiting to be transferred by 

hand or sling to a processing station on board. Net pens measuring three feet wide, six 

feet long, and three feet deep can safely hold about 20 adult shortnose sturgeon or 

comparably sized juvenile Atlantic, Gulf or green sturgeon when temperatures are below 
13 

15°C (Doug Peterson, University of Georgia, pers. comm.). Larger net pens (8 feet long) 

are required for holding adult Atlantic, green, and Gulf sturgeon or they should be 

processed as quickly as possible (or scheduled first) instead of subjected to confined 

holding conditions. When water temperature is between 15° and 25°C, fewer fish should 

be held in the same enclosure because overcrowding animals amplifies short term stress, 

particularly at higher temperatures (Safi et al. 2006). If the fish are being held on-board a 

vessel in a holding tank, compressed oxygen should be added to increase DO in the 

water. If the researcher observes a visually stressed sturgeon, efforts should be made to 

revive the fish and release it in a healthy condition. In some cases, recovery can be 

achieved by allowing a sturgeon to rest in an appropriately sized net pen for several hours 

prior to release. 

Sturgeon should never be held in gillnets if there isn’t enough room to safely hold them 

in net pens. In some rivers with large populations of sturgeon, catches can exceed the 

number of fish that can possibly be held safely in live cars or net pens. In such cases, 

researchers should have multiple holding bins at their disposal. If more fish are captured 

than can be processed and released within two hours, those excess fish may need to be 

released to minimize stress or lethal injury. 

When sturgeon are held on-board research vessels, they should be placed in flow through 

tanks where the total volume of water is replaced every 15 to 20 minutes. Traditionally, 

some species of sturgeon have been held for research purposes by tethering with ropes 

looped around tails to the sides of research vessels until they can be handled. In a study 

of lake sturgeon (Axelsen and Mauger 1993 cited in Dick et al. 2006), tethered fish 

experienced greater stress and higher mortality than sturgeon kept in uncrowded cages. 

Therefore, NMFS recommends only using on-board holding tanks or net pens large 

enough to hold a large sturgeon. NMFS does not recommend holding any sturgeon by 

tethering its caudal peduncle to the research vessel. However, while a rope should never 

be tied around the caudal peduncle, it may be necessary to use a rope placed under the 

sturgeon immediately posterior to the pectoral fins when moving large sturgeon from net 

pens onto the boat. 

Following handling procedures, fish should be returned to the net pen for observation and 

to ensure full recovery prior to release. Total holding time in the net pens would be 

variable depending on water temperature and the condition of each fish, however, the 

maximum amount of time a fish should be held after removal from capture gear is 

approximately two hours, unless more time is needed to recover from the effects of an 

anesthetic or because prolonged holding would benefit a sturgeon. When water 

temperature is above 25°C for Gulf, shortnose, and Atlantic sturgeon, or 23°C for green 

sturgeon, they should be held for as little time as possible. Holding time includes the 

time to remove any other captured sturgeon, time to process other fish, and time 

necessary for recovery ensuring the safety of the fish. 

Prior to release, sturgeon should be examined and, if necessary, recovered by holding fish 

upright and immersed in river water, gently moving the fish front to back, aiding 

freshwater passage over the gills to stimulate it. The fish should be released when 
14 

showing signs of vigor and able to swim away under its own power. A spotter should 

watch the fish, making sure it stays submerged and does not need additional recovery. 

Recommendations 

Proper Handling of Sturgeon 

• NMFS strongly recommends standard handling procedures performed on all 

sturgeon captured including measuring, weighing, PIT tagging, and tissue 

sampling. 

• Only one additional discretionary procedure to the standard handling procedures 

should be performed on each sturgeon, thus minimizing handling time prior to 

release. 

• When water temperatures are above 23°C for green sturgeon or 25°C for Gulf, 

shortnose, or Atlantic sturgeon, the extent of research should be limited to the 

standard handling procedures of measuring, weighing, PIT tagging, and tissue 

sampling. 

• During handling procedures, each fish should be immersed in a continuous stream 

of ambient water passing over the sturgeon’s gills. 

• Researchers should attempt to support larger sturgeon in slings preventing 

struggle during transfer. 

• If the researcher observes a severely stressed sturgeon, efforts should be made to 

revive the fish and release it in a healthy condition. 

Short-Term Holding 

• Sturgeon should never be held in gillnets while waiting to be handled, but should 

instead be transferred to a net pen for holding. 

• NMFS recommends only using on-board holding tanks or net pens large enough 

to hold a large sturgeon. NMFS does not recommend tethering sturgeon to the 

boat by its caudal peduncle. 

• The maximum amount of time a fish should be held after removal from capture 

gear is approximately two hours, unless more time is needed to recover from the 

effects of an anesthetic or because prolonged holding would benefit a sturgeon. 

• Adult Atlantic, green, and Gulf sturgeon over six feet in length should be 

processed as quickly as possible (or scheduled first) instead of subjected to 

confined holding conditions. 
15 

Standard Research Methods 

Upon capturing a green, Gulf, shortnose, or Atlantic sturgeon, there are several research 

procedures strongly recommended on all sturgeon. First, the captured fish is to be 

measured. The sturgeon should also be weighed if possible. It can also be photographed, 

if possible. Then, their entire bodies should be scanned for previously inserted PIT tags; 

and, if none are found, one should be properly inserted. Finally, a small sample of the 

soft tissue of the pelvic fin should be removed for genetic identification. 

Measuring 

Standardized length measurements for all sturgeon should be taken from the snout to the 

fork in the tail (i.e., fork length – FL). The measuring device should be a solid ruler or 

board, so the measurement does not measure the curvature of the body. Additional length 

measurements should be taken at the researcher’s discretion for total length (TL) or head 

length (Figure 1). While the heterocercal tail of larger fish may be damaged or 

shortened, the total length can still be obtained by pressing down the tail at the caudal 

peduncle and measuring to the tip of the tail. Girth measurements should also be taken at 

the widest part of the body. While not mandatory, measurements of the ratio of mouth 

width to interorbital width can also be obtained to differentiate between shortnose and 

Atlantic sturgeon (Dadswell et al. 1984). Interorbital width is measured as the distance 

between the lateral margins of the bony skull at the midpoint of the orbit and mouth 

width is measured as the distance between the left and right inside corners of the closed 

mouth (i.e., excluding the lips) (Figure 1). 

Figure 1. Diagram of different types of measurements for sturgeons. Drawings by Eric 

Hilton, Virginia Institute of Marine Science. 
16 

Weighing 

All captured sturgeon should be weighed if possible. Weights allow a better 

understanding of the conditioning of captured sturgeon during various seasons of the year 

or life span of the fish. For weighing sturgeon, animals should be supported with a sling 

or net and handling should be minimized throughout the procedure. 

Boats used for researching green, Gulf, and Atlantic sturgeon should accommodate larger 

fish with scales available to safely weigh a 200 pound fish. When targeting shortnose 

sturgeon (or juvenile green, Gulf, or Atlantic sturgeon), hand-held sling scales are 

acceptable. When using a bench scale or platform scale to weigh large sturgeon, a five to 

six foot flat platform will be necessary to support the fish. 

Photographing 

When handling sturgeon, optional photography is often used to document the health of 

fish, research methods, and any identifying marks on the sturgeon potentially useful in 

the future. Although it is recommended to take as many pictures as needed, researchers 

should do so without interfering with other research activities. 

PIT Tags 

Every sturgeon should be scanned for PIT tags along its entire body surface ensuring it 

has not been previously tagged. Untagged sturgeon should then be appropriately PIT 

tagged (Figure 2) and the identifying number recorded. Each PIT tag consists of 

integrated circuitry and an antenna encapsulated in glass. PIT tags are “passive” because 

they contain no batteries; their internal code is activated and transmitted to the receiver 

when exposed to the transceiver’s electromagnetic signal. The newest PIT tags, and 

those recommended by NMFS, use a frequency of 134.2 kHz. 

Standardized PIT tag placement for Gulf, green, Atlantic, and shortnose sturgeon would 

enable subsequent researchers to locate prior PIT tags quickly and consistently. 

Sturgeon, are large fish growing a considerable amount from the time they’re first PITtagged until they reach their adult size. If muscles grow over the PIT tag as they mature, 

the tag can become increasingly more difficult to read. 

For this reason, NMFS strongly recommends PIT tag placement in all four sturgeon 

species to be located to the left of the spine, immediately anterior to the dorsal fin, and 

posterior to the dorsal scutes (Figure 2). This positioning would optimize PIT tag 

readability over the animal’s lifetime as sturgeon experience the least new muscle growth 

in this location during their lifetimes (Berg 2004, Simpson and Fox 2006). After the tag 

is inserted, it should be scanned to ensure it is readable before the fish is released. If 

necessary, to ensure tag retention and prevent harm or mortality to small juvenile 

sturgeon of all species, the PIT tag can also be inserted at the widest dorsal position just 

to the left of the 4th dorsal scute. 
17 

Figure 2. Standardized location for PIT tagging all green, Gulf, 

Atlantic, and shortnose sturgeon. (Photo by James Henne, USFWS) 

PIT tags have the highest reported retention rate of all identification tags, though they are 

not visible to the researcher or fisherman upon capture. Clugston (1996) found PIT tags 

implanted in gulf sturgeon have approximately a 90% retention rate. Musick and Hager 

(2007) tagging 445 Atlantic sturgeon reported a 99% retention rate of PIT tags after 96 

hours. Smith et al. (1990) noted 100% retention after 60 days in wild shortnose sturgeon. 

In the Penobscot River, retention rates for PIT tags in Atlantic sturgeon were 93% after as 

much as 8.8 years (Gayle Zydlewski, University of Maine, pers. comm.). Nelson et al.

(2007) report approximately 100% retention of PIT tags in recaptured white sturgeon. 

Other researchers have had different results. Researchers with EDI Environmental 

Dynamics (2006) reported recapturing three white sturgeon, with 66% retention of PIT 

tags. DeHaan et al. (2008) recorded 51 to 95% retention when PIT-tagging juvenile 

pallid sturgeon, which is similar to rates observed by Henne et al. (unpublished). 

As with all research procedures, there is a risk of injury or mortality either directly or 

indirectly related to PIT tagging. When PIT tags are inserted into animals having large 

body sizes relative to tag size, empirical studies generally conclude they have no adverse 

effect on the growth, survival, reproductive success, or behavior of individual animals 

(Brännäs et al. 1994, Elbin and Burger 1994, Keck 1994, Jemison et al. 1995, Clugston 

1996, Skalski et al. 1998, Hockersmith et al. 2003). However, smaller sturgeon may 

experience mortality within the first 24 hours, usually as a result of inserting the tags too 

deeply or from pathogenic infection. When analyzing mortality of small sturgeon caused 

by PIT tags, Henne et al. (2008) found 11 and 14 mm tags inserted into shortnose 

sturgeon longer than 300 mm was safe. In this study, they found that when fish are under 

300 mm, factors other than length, such as weight or condition, most influence the 

likelihood of mortality. Therefore, NMFS recommends only sturgeon over 300 mm 

should receive PIT tags. 

A negative aspect of using PIT tags in sturgeon research is the difficulty for NOAA 

observers or non-researchers to detect tags in recaptured sturgeon without the benefit of a 

PIT tag reader. Rien et al. (1994) and Nelson et al. (2004) recommend removal of the 

second left lateral scute indicating the presence of a PIT tag in white sturgeon. This 

methodology has been subsequently used for green sturgeon as well. While removal of 
18 

scutes rarely results in bleeding, and is not considered deleterious, there are other, safer 

means for externally marking sturgeon. NMFS believes a standardized PIT tag location 

is less stressful to animals and is easily located. If an external mark is necessary, NMFS 

recommends using other external tags identified in this document. Those external tags 

are not only obvious to other researchers, but also to the general public for identifying 

recaptured animals to alert researchers of their recapture. NMFS therefore recommends 

using external tags to identify the presence of a PIT tag, if necessary, but researchers 

should not remove scutes from sturgeon for any reason. 

Genetic Tissue Sampling 

Tissue sampling is a common practice in fisheries science characterizing the genetic 

“uniqueness” and quantifying the level of genetic diversity within a population. 

NMFS strongly recommends genetic tissue samples be taken from every sturgeon 

captured unless, due to marks or tags, the researcher knows a genetic sample has already 

been obtained. Tissue samples should be a small (1.0 cm2

) fin-clip collected from soft 

pelvic fin tissues using a pair of sharp scissors. Tissue samples should be preserved in 

individually labeled vials containing 95% ethanol. There is no evidence that this 

procedure harms any species of sturgeon. 

Recommendations 

Strongly Recommended

• Researchers should measure all captured green, Gulf, Atlantic, and shortnose 

sturgeon. The sturgeon should also be weighed, if possible. 

• Researchers should scan captured sturgeon for previously inserted PIT tags; and, 

if none are found, one should be properly inserted. 

• Researchers should remove a small tissue sample by clipping the soft tissue of the 

pelvic fin. 

Measuring 

• Standardized length measurements for all sturgeon should be taken from the snout 

to the fork in the tail. 

• NMFS recommends measuring the ratio of mouth width to interorbital width to 

differentiate shortnose and Atlantic sturgeon. 

PIT Tags 

• NMFS recommends PIT tag placement in all four sturgeon species to be located 

to the left of the spine, immediately anterior to the dorsal fin, and posterior to the 

dorsal scutes. 

• NMFS recommends using 134.2 kHz PIT tags. 

• If necessary, to ensure tag retention and prevent harm or mortality to small 

juvenile sturgeon of all species, the PIT tag can also be inserted at the widest 

dorsal position just to the left of the 4th dorsal scute. 

• NMFS recommends only sturgeon over 300 mm should receive PIT tags. 

• NMFS recommends using external tags to identify the presence of a PIT tag, if 

necessary, but researchers should not remove scutes from sturgeon for any reason. 
19 

Genetic Tissue Sampling 

• NMFS strongly recommends genetic tissue samples be taken from every sturgeon 

captured unless, due to marks or tags, the researcher knows a genetic sample has 

already been obtained. 

• Tissue samples from Gulf, green, Atlantic, and shortnose sturgeon should be 

archived at the NOAA/NOS Tissue Archive in Charleston, South Carolina. 

Proper certification, identity, and chain of custody of samples should be 

maintained during transfer of tissue samples. 
20 

Anesthetization 

Anesthetics are physical or chemical agents preventing the initiation and conduction of 

nerve impulses (Summerfelt and Smith 1990). Therefore, the primary functions of 

anesthetics on ESA listed sturgeon are to immobilize the animal allowing precise, 

autorized procedures to be performed while blocking nerve impulses which might 

otherwise adversely affect the fish. This section, therefore, attempts to balance the risk of 

stress from invasive procedures with the risk posed by using an anesthetic, while also 

considering the risk of an unanesthetized sturgeon moving suddenly during a procedure 

resulting in trauma or hemorrhaging. 

Invasive research activities can be stressful to fish, even if immobilized. The use of an 

anesthetic reduces the potential for short term stress response and risk of mortality during 

those procedures (Iwama et al. 1989, Small 2003, Wagner et al. 2003, Coyle et al. 2004, 

Roubach et al. 2005, Wanner et al. 2007). However, the use of some anesthetics have 

also proven to be stressors to fish (Iwama et al. 1989) as evidenced by the buildup of the 

cortisol hormone. NMFS recommends that noticeably stressed sturgeon should not be 

anesthetized. 

Documented lethal or sub-lethal effects caused by improper dosage or exposure of 

anesthetics (Iwama et al. 1989, Summerfelt and Smith 1990) raises concerns whether it is 

acceptable to use anesthetic when handling listed Gulf, green, shortnose, or Atlantic 

sturgeon. In tests where anesthetics were not used during invasive procedures, cortisol 

levels were found significantly higher than when fish were anesthetized with tricaine 

methanesulfonate (MS-222) or clove oil (Wagner et al. 2003). Conversely, Wagner et al.

(2003) found unanesthetized fish had lower cortisol levels than either of two anesthetized 

groups after one hour, demonstrating recovery of fish is more rapid without 

anesthetization. Nevertheless, in controlled studies when prolonged handling took place 

(30 minutes or more), Strange and Schreck (1978) documented fish had a higher survival 

rate when anesthetized. 

Summerfelt and Smith (1990) and Bowser (2001) note a normal condition and six stages 

of anesthesia: light sedation, deep sedation, partial loss of equilibrium, total loss of 

equilibrium, loss of reflex reactivity, and asphyxia (Table 4). Light sedation occurs when 

there is a slight loss of reactivity, while deep sedation occurs when only the strongest 

external stimuli will elicit a response, but in both cases, the fish is able to maintain 

equilibrium. Partial loss of equilibrium is also characterized by partial loss of muscle 

tone and an increase in opercular movement, while total loss of equilibrium is 

characterized by total loss of muscle tone, the loss of spinal reflexes, and slow and steady 

opercular rate. The loss of reflex reactivity is when the fish losses all reflex response, but 

also when the heart rate becomes very slow and the opercular movements become slow 

and irregular. The final stage of anesthesia is a complete medullary collapse, when 

opercular movement ceases. Death is typically caused by an overdose or overexposure 

leading to eventual mortality. 
21 

Table 4. Stages of anesthesia (Summerfelt and Smith 1990). 

Stage Descriptor Behavioral Response of Fish 

0 Normal Reactive to external stimuli; opercular rate and 

muscle tone normal 

I Light sedation Slight loss of reactivity to external stimuli; 

opercular rate slightly decreased; equilibrium 

normal 

II Deep sedation Total loss of reactivity to all but strong external 

stimuli; slight decrease in opercular rate; 

equilibrium normal 

III Partial loss of equilibrium Partial loss of muscle tone; swimming erratic; 

increased opercular rate; reactivity only to strong 

tactile and vibration stimuli 

IV Total loss of equilibrium Total loss of muscle tone and equilibrium; slow but 

regular opercular rate; loss of spinal reflexes 

V Loss of reflex reactivity Total loss of reactivity; opercular movements slow 

and irregular; heart rate very slow; loss of all 

reflexes 

VI Medullary collapse (asphyxia) Opercular movements cease; cardiac arrest usually 

follows quickly 

The primary risks associated with anesthetizing sturgeon are overexposure and 

overdosing. Overexposure can occur when sturgeon are left in an anesthetic bath longer 

than necessary to achieve narcosis. Fish often have difficulty recovering with normal 

response time when overexposed, and sometimes will not respond for extended periods 

requiring continuous respiration to revive them. Overdosing can take place when the 

concentration of anesthetic is higher or more toxic than fish can tolerate. Both conditions 

often result in immediate or delayed mortality. As an anesthetic is applied, the sturgeon’s 

opercular movement should be monitored closely. It should not be allowed to stop as this 

condition could result in blood hypoxia and high stress response, or even mortality of the 

anesthetized animal (Iwama et al. 1989). 

There are various research activities commonly performed on sturgeon that present 

enough risk to the fish that they should only be done using anesthesia (Table 5). 

However, the same level of narcosis is not needed for each activity and therefore the 

researcher would not use the same concentrations of anesthetic. Physical restraint is not 

an appropriate substitute for anesthetization. 

The rate at which anesthesia is induced in a fish is also important at minimizing stress. 

Prolonged induction generally leads to increased stress responses (e.g. prolonged 

thrashing during excited phase), while excessively rapid induction times (<1 minute) 

risks taking the fish beyond the surgical anesthesia plane because animals may skip 

typical behavioral signs characterizing stages of anesthesia. NMFS recommends 

initiating anesthesia gradually to reduce the risks of overdosing. NMFS also 

recommends monitoring the sturgeon during induction to avoid overexposure. If the 

desired stage of narcosis cannot be reached within 15 minutes (Summerfelt and Smith 

1990), the sturgeon should be placed in freshwater to recover before being released. 
22 

Table 5: Procedures and stages of anesthesia. 

Procedure Stage of Anesthesia (see Table 4) 

Internal tagging III 

Biopsy III 

Laparoscopy IV 

Gastric lavage I 

Boroscope 0 or I 

Fin ray sectioning II 

Genetic fin clip 0 

Blood sample 0 

PIT tag 0 

External tagging 0 but I is acceptable if necessary 

Cold water species respond more rapidly and at lower doses to chemical anesthetics than 

do warm water species (Bowman et al. 2003, Coyle et al. 2004). Currently, it has not 

been demonstrated if shortnose, Atlantic, Gulf, or green sturgeon exhibit variable interor intra-species responses to chemical anesthetics with respect to temperature. As 

identified previously, however, larger green sturgeon grow more optimally at cooler 

temperatures than do shortnose or Atlantic sturgeon. This suggests green sturgeon are 

better adapted to cooler waters, may also be more likely to respond to lower levels of 

anesthetic than shortnose or Atlantic sturgeon. Correspondingly, Gulf sturgeon may need 

higher doses than the other species at cooler temperatures. Likewise, northern 

populations of shortnose and Atlantic sturgeon may be better adapted to cooler waters 

and respond differently to anesthesia. 

Chemical Anesthetic 

MS-222

A wide variety of chemical compounds have been utilized to anesthetize fish in fisheries 

research. However, tricaine methanesulfonate (MS-222) is the only anesthetic with a 

label for use with fish granted by the Food and Drug Administration (FDA) and as such, 

is the only chemical anesthetic recommended by NMFS for use on green, Gulf, Atlantic, 

and shortnose sturgeon. 

MS-222 is absorbed rapidly through the gills and it prevents the generation and 

conduction of nerve impulses, with direct actions on the central nervous system and 

cardiovascular system. MS-222 is excreted in fish urine within 24 hours and tissue levels 

decline to near zero in the same amount of time (Coyle et al. 2004). 

Proper dosing depends on the degree of anesthetization desired, the species and size of 

fish, water temperature and water hardness. In general, levels of MS-222 recommended 

do not typically exceed 100mg/L for salmonids or 250 mg/L for warm water fish (Coyle 

et al. 2004). To euthanize fish using MS-222, the recommended dosage varies from 150 

to 500 mg/L for one minute or more depending on the species (DeTolla et al. 1995, Cho 

and Heath 2000, Callahan and Noga 2002, Borski and Hodson 2003). 
23 

There are two methods commonly used by sturgeon researchers to anesthetize sturgeon. 

The first method incorporates a “knockout” initiation dose of MS-222 followed by a safer 

maintenance concentration (DeTolla et al. 1995, Callahan and Noga 2002, Thorsteinsson 

2002, Borski and Hodson 2003). Alternately, researchers anesthetize sturgeon using the 

lowest possible dose of MS-222, raising it to achieve the desired stage of narcosis based 

on the procedure (Table 5). Neither method, when performed correctly, is safer than the 

other. However, more risk is associated with overdosing fish exposed to higher induction 

rates. 

For most procedures, sturgeon should initially be lightly anesthetized with MS-222, and 

if needed, more should be added only to the level considered necessary to perform the 

appropriate procedures. MS-222 solutions are highly acidic, therefore the pH of the 

solution should be buffered to a neutral pH with equal amounts of sodium bicarbonate 

prior to use. In cooler water temperatures, either higher doses or longer exposure times 

may be necessary to achieve the proper narcosis because the absorption rate is lower at 

lower temperatures (Coyle et al. 2004). Additionally, because MS-222 is a hypoxic 

agent, the anesthetic container should be vigorously aerated to maintain DO levels 

equivalent to ambient river water. 

Total loss of equilibrium (Stage IV) is the deepest level of narcosis acceptable for 

anesthetizing listed sturgeon. It may not be possible to reach this stage of narcosis by 

gradually increasing the dosage and instead, the researcher would need to begin with a 

high induction dose and then drop back to a maintenance dose. Because of the risks 

associated with this type of anesthetization, NMFS recommends inexperienced 

researchers first conduct this type of anesthesia in a laboratory using a heart rate monitor 

to prevent overdose. Only once a researcher has demonstrated the ability to consistently 

perform this type of anesthetization safely should they do this in the field. 

When immersed in MS-222, sturgeon will initially experience rapid gill movement 

followed by marked reduced gill movement as the agent begins to have an effect. As gill 

movement slows, sturgeon will lose equilibrium and eventually turn upside down or float 

to the surface. At this stage, sturgeon should be watched closely to confirm continuous 

involuntary gill movement. If the procedure is brief, once the desired stage of anesthesia 

has been reached, sturgeon may be placed on a surgical cradle and the gills irrigated with 

fresh water to ensure respiration and to begin recovery as the procedure is quickly 

completed. After completing the procedure, the fish should be placed in a clean, 

anesthetic free recovery tank and observed until fully recovered. Once recovered, the 

sturgeon can be released. 

Following is a review of the various concentrations and induction methods of MS-222 

when anesthetizing Gulf, Atlantic, shortnose, and green sturgeon. Fleming et al. (2003a) 

suggested concentrations of MS-222 of up to 400 mg/L failed to adequately anesthetize 

Gulf sturgeon. These researchers concluded the anesthetic was potentially dangerous to 

the sturgeon. However, Hernandez-Divers et al. (2004) successfully anesthetized Gulf 

sturgeon submerging them in an initiating dose of 250 mg/L followed by a maintenance 

bath of 87.5 mg/L. Harris et al. (2005) anesthetized Gulf sturgeon using 160 mg/L MS-
24 

222. Parkyn et al. (2006) anesthetized Gulf sturgeon using a single phase induction of 

150 mg/L MS-222. Lankford et al. (2005) anesthetized green sturgeon placing them in 

concentrated baths of 350 mg/L of MS-222 followed by less concentrated doses of 150 

mg/L. Kaufman et al. (2007) anesthetized green sturgeon using 350 mg/L removing 

them from the solution when anesthetized. However, Serge Doroshov, (University of 

California Davis, pers. comm.) regularly uses 100 mg/L when working on green 

sturgeon. Joe Cech (University of California Davis, pers. comm.) starts green sturgeon 

anesthesia in baths of 150 mg/L and then when respiration stops, places them in a second, 

less concentrated bath of 75 mg/L. The majority of shortnose and Atlantic sturgeon 

researchers interviewed for this document reported concentrations of MS-222 from 50 to 

100 mg/L were sufficient to induce anesthesia for most invasive procedures (Boyd 

Kynard Permit #1549, Mark Collins Permit #1447, Michael Kennison Permit #1595, 

Doug Peterson Permit #10037, Haley 1998, Oakley and Hightower 2007, Savoy 2007). 

The USFWS’ Biological Procedures and Protocols for Researchers and Managers 

Handling Pallid Sturgeon recommends using MS-222 at doses between 50 and 150 mg/L 

(USFWS 2008). 

Induction and recovery times for chemical anesthetics vary based on the dosage level and 

duration the fish is under anesthesia. For rainbow trout in MS-222, Wagner et al. (2003) 

found induction takes two to three minutes at 60 mg/L with recovery taking 5 to 6 

minutes. For Gulf sturgeon in MS-222, Hernandez-Divers et al. (2004), when initiating 

anesthesia at 250 mg/L, induction took 5 to 11 minutes before lowering the dosage to 

87.5 mg/L, after which recovery took 3 to 13 minutes. For green sturgeon at 50 to 100 

mg/L MS-222, induction and recovery both required 10 to 15 minutes at 18° to 21°C, but 

at cooler temperatures it took longer (Joel Van Eenannaam, University of California 

Davis, pers. comm.). 

Sturgeon face several risks posed by MS-222, such as overdose, increased stress, or being 

released prior to recovering. Weakened fish are more susceptible to anesthetic shock and 

thus are more likely to be accidentally overdosed (Coyle et al. 2004). Even when 

anesthetized with MS-222, fish still experience elevated levels of plasma cortisol, 

indicating they are stressed either by handling or by additive stress of MS-222 (Coyle et 

al. 2004). After being handled under anesthesia, plasma cortisol levels increased 8 times 

over base in channel catfish (Small 2003) and nine times over base in rainbow trout 

(Wagner et al. 2003). Studies by Pirhonen and Schreck (2003) found fish anesthetized 

with MS-222 ate significantly less (15-20%) than control fish. If the dose of MS-222 is 

too high or the exposure is too long, recovery is longer if it occurs at all. Therefore, 

NMFS recommends monitoring sturgeon closely during recovery and taking protective 

measures if fish appear stressed and not recovering normally (e.g., providing 

supplementary DO and moving water across the gills until fully recovered). 

Recovery is also influenced by the size and sexual condition of fish. Because MS-222 is 

fat soluble (Coyle et al. 2004) longer recovery times are experienced by larger sturgeon 

and gravid females. Holcomb et al. (2004) showed doses of 225 mg/L MS-222 had no 

effect to eggs or sperm of white sturgeon and could be used to harvest gametes. 

However, doses of 2,250 mg/L resulted in lower hatching success and doses of 22,500 
25 

mg/L resulted in complete loss of fertility. At the dosages typically used by researchers 

to anesthetize sturgeon, however, no impact to their eggs is expected. 

Although the FDA permits the use of MS-222, it also requires a 21 day withdrawal period 

before an anesthetized fish can be consumed. This poses concerns for humans when non 

listed fish are released into the wild where they may be consumed. However, a 21 day 

withdrawal is not a consideration for threatened or endangered sturgeon, as taking or 

possessing them is prohibited by the ESA. Therefore, no external marks or tags are 

required for Gulf, green, Atlantic, or shortnose sturgeon following anesthetization with 

MS-222. 

Clove Oil

Clove oil is approximately 90 to 95% eugenol with smaller portions of methyleugenol 

and isoeugenol and was initially experimented with as a substitute for MS-222 (Bowman 

et al. 2003). Showing promise as an anesthetic, it was marketed as AQUI-S (isoeugenol , 

2-methoxy-4-propenylphenol) in an attempt to gain FDA approval. However, in 2007, 

the National Toxicology Program concluded exposure of male mice to isoeugenol 

resulted in clear evidence of cancer. As a result of its concern that isoeugenol’s 

carcinogenic properties could be transmitted through the food web, the FDA’s Center for 

Veterinary Medicine officially rescinded authorization for the “investigational food use” 

of AQUI-S under INAD 10-541 (AADAP 2008). Consequently, both NMFS and the 

FDA (2007) are concerned isoeugenol could have direct adverse effects to threatened and 

endangered aquatic species. NMFS does not authorize the use of clove oil or AQUI-S on 

Atlantic, green, Gulf, or shortnose sturgeon. 

Physical Anesthetic 

Electronarcosis

Electronarcosis (also referred to as electroanesthesia and galvanonarcosis) is a nonchemical method of anesthetization and, as such, does not require FDA approval. 

Researchers investigating the use of electricity to immobilize fish have used various 

methods and species of fishes. Alternating current (AC), rectified AC, constant direct 

current (CDC), and pulsed direct current (PDC) have all been tested (Hartley 1967, 

Walker et al. 1994, Barton and Dwyer 1997, Henyey et al. 2002). Some researchers 

leave the electricity on for the entire time the fish is immobilized (Gunstrom and Bethers 

1985) while others apply a short burst of relatively high voltage resulting in 

immobization of the fish for several minutes after the electric current is discontinued 

(Sterritt et al. 1994). Much of what has been learned about electronarcosis is based on 

the same principles applied during electrofishing. 

Fish exposed to electric current may show electrotaxis (forced swimming), electrotetanus 

(muscle contractions), or electronarcosis (muscle relaxation). AC causes tetanus (Henyey 

et al. 2002) and at higher voltages pulsed direct current causes tetanus, whereas constant 

direct current causes narcosis first, and then will eventually cause tetanus as the voltage is 

increased (Summerfelt and Smith, 1990). Typically, when researchers have studied 

electronarcosis, the electricity used was either AC or PDC, or was CDC of a sufficiently 
26 

high voltage that the fish were immobilized by electrotetanus. Further, most studies 

using AC and PDC reported adverse effects including some bruising, burning, 

hemorrhaging, and mortality (Tipping and Gilhuly 1996, Redman et al. 1998, Holliman 

and Reynolds 2002). Consequently, NMFS does not recommend using AC or PDC 

currents for inducing anesthesia in listed sturgeon. When using CDC, the risks to 

sturgeon are over-applying the direct current resulting in either tetany or cessation of 

opercular movement. These adverse affects can be avoided by monitoring the sturgeon 

and reducing the voltage depending on the fish’s behavior. 

Henyey et al. (2002) describe using low voltage CDC to induce electronarcosis (muscle 

relaxation) in shortnose sturgeon without any changes in swimming or feeding behavior, 

burns, bruising, or mortality after monitoring the fish for six weeks (Boyd Kynard, 

USGS, pers. comm.). All evidence indicates electronarcosis induced by the method 

described is similar to the condition induced by chemical anesthetics; nevertheless, more 

research is needed on the physiological mechanisms by which it works. NMFS 

recommends low voltage direct current electronarcosis as described by Henyey et al.

(2002) as a viable alternative to chemical anesthesia. 

Electronarcosis has been used successfully by Boyd Kynard (USGS, pers. comm.) to 

anesthtize shortnose sturgeon since the 1980s. Since 2004, USFWS researchers in 

Maryland have also followed the Henyey et al. (2002) protocol to anesthetize Atlantic 

and shortnose sturgeon on the Potomac River and Chesapeake Bay with no adverse 

affects reported (Mike Mangold, USFWS, pers. comm.). Researchers in South America 

have also followed these methods reporting similar success (Alves et al. 2007). 

As described in Henyey et al. (2002), a tank is prepared by positioning positive cathode 

and negative anode plates at opposite ends. With the sturgeon oriented head towards the 

cathode, a CDC is applied quickly so the fish loses equilibrium and then the voltage is 

adjusted downward until the fish is relaxed and exhibiting strong opercula movement. In 

practice, when inducing electronarcosis, if gill ventilation becomes irregular or stops, the 

electric current should be decreased and the fish will recover steady ventilation 

immediately (Boyd Kynard, USGS, pers. comm.). The amperes should be set to the 

minimal level (0.01A). Depending on the individual sturgeon and water chemistry, about 

0.3 to 0.5 volts per centimeter is recommended to immobilize sturgeon. Typically, 

sturgeon should be supported by a net so only half of the body either dorsal or ventral 

depending on the work being conducted, is out of the water. Under these conditions, the 

researcher will feel nothing while working in the water (Hartley 1967, Boyd Kynard, 

USGS, pers. comm.) but researchers with sensitive skin or hand abrasions are also 

encouraged to wear rubber gloves during the procedure. 

Induction and recovery from electronarcosis both require less than 10 seconds because as 

soon as fish are placed in or removed from the electrical current, it is no longer 

anesthetized (Gunstrom and Bethers 1985, Summerfelt and Smith 1990, Henyey et al.

2002). Henyey et al. (2002) state electronarcosis is ideal for non-invasive research. The 

methods in Henyey et al. (2002) elicited narcosis, not tetany; and Boyd Kynard (USGS, 

pers. comm.) states narcosis is induced by blocking nerve impulses at the medulla 
27 

oblongata. Kynard and Lonsdale (1975) demonstrated electronarcosis and MS-222 

yielded similar states of muscle relaxation and immobility. 

Recommendations 

General 

• NMFS recommends that noticeably stressed sturgeon should not be anesthetized. 

• Physical restraint is not an appropriate substitute for anesthetization in procedures 

requiring anesthesia. 

• NMFS recommends initiating both chemical and physical anesthesia gradually to 

reduce the risks of overdosing. 

Chemical Anesthetic 

• Because of the risks associated with high initial induction doses followed by a 

lower maintenance dose of MS-222, NMFS recommends using this technique in a 

controlled environment such as a laboratory and also using a heart rate monitor to 

prevent overdosing. 

• NMFS also recommends monitoring the sturgeon during induction to avoid 

overexposure and if the desired stage of narcosis cannot be reached within 15 

minutes, the sturgeon should be placed in freshwater to recover before being 

released. 

• Tricaine methanesulfonate (MS-222) is the only chemical anesthetic with a label 

for use on fish granted by the FDA and as such, is the only chemical anesthetic 

recommended by NMFS for use on green, Gulf, Atlantic, and shortnose sturgeon. 

Dosages of MS-222 should be between 50 and 150 mg/L as identified in the pallid 

sturgeon protocols (USFWS 2008) and by green, Gulf, Atlantic, and shortnose 

sturgeon researchers. 

• NMFS recommends monitoring sturgeon closely during recovery and taking 

protective measures if fish appear stressed and not recovering normally (e.g., 

providing supplementary DO and moving water across the gills until fully 

recovered). 

• A 21 day withdrawal, normally associated with the use of MS-222 on food fish, is 

not a consideration for threatened or endangered sturgeon, as taking or possessing 

them is prohibited by the ESA. 

• NMFS does not authorize the use of clove oil and AQUI-S on Atlantic, green 

Gulf, or shortnose sturgeon. 

Physical Anesthetic 

• NMFS recommends low voltage direct current electronarcosis as described by 

Henyey et al. (2002) as a viable alternative to chemical anesthesia but does not 

recommend using AC or PDC currents for inducing anesthesia in listed sturgeon. 
28 

Tagging 

Tagging is an essential function of sturgeon research, serving to identify unique 

information about a captured or recaptured animal. PIT tags, as discussed earlier, should 

be inserted in all Gulf, green, shortnose, and Atlantic sturgeon without a PIT tag. 

Determining the life history, morphology, behavior, movement, and physiology of 

sturgeons are all highly dependent on proper tagging methods. Because sturgeon can live 

for decades, it is essential tags be retained for extended periods. In addition, because 

sturgeon exhibit very rapid juvenile growth rates and can achieve very large sizes, tags 

must be retained even as the tag placement area changes size and shape. Moreover, 

sturgeon are adept at shedding external tags and can also extrude internal tags through the 

body wall (Kieffer and Kynard 1993). Consequently, sturgeon researchers should keep 

well informed on the effectiveness of tagging methods and the technology best suited for 

local conditions. 

Tagging varies based on tag function, location, method, technology, retention rates, and 

size. Internal tags (acoustic or radio) are surgically implanted in sturgeon for tracking 

movements, whereas externally mounted tags can be used for tracking or identification. 

Despite lower retention rates for some external tags, there are situations where external 

tags are the only option, such as tracking pre-spawning females. External archival tags 

and satellite tags can also passively record water quality information or geographic 

position without arrays. Other types of external-identifier tags are useful when nonresearchers are involved in research activities, such as studies relying on fishermen to 

return data from tags on marked fish. 

Telemetry Tags 

Acoustic tags outperform radio tags in deeper water (or saline water) where sturgeon 

spend a majority of their lives; however, acoustic tags have disadvantages associated with 

limited range and ineffectiveness in turbulent or turbid waters. Acoustic signals can be 

monitored by field crews using either mobile hydrophones or, more commonly, 

stationary hydrophone arrays. Because the stationary arrays are designed to passively 

capture the location of transmitted signals from near-by fish, many researchers are 

converting to acoustic tag technology, collecting data over a longer period of time and 

downloading it at later intervals (Reine 2005). 

Radio transponders emit radio signals from transmitter antennae to the atmosphere where 

they can then be monitored by researchers with a receiving antenna. For highly 

migratory species such as sturgeon, researchers can locate and track fish at distances up 

to three kilometers via airplane. Radio signals are also effective in environments having 

more physical disruptions such as turbidity (Thorsteinsson 2002). Combined acoustic 

and radio transmitter (CART) tags provide the researcher the advantages of each 

transmitter type. 

Implanting internal telemetry tags is stressful to sturgeon and should be done using 

anesthesia. To gain access to the abdominal cavity, a two to four centimeter incision is 

made between the 3rd and 4th ventral scute between the anal and pelvic fin slightly left or 
29 

right of the mid-ventral line. Internal tags should be coated with a biologically inert 

substance, soaked in alcohol and allowed to dry, and then pushed deeply into the 

abdominal cavity to prevent tags from rubbing against the incision (Kynard and Kieffer 

1991). In studies by Kynard and Kieffer (1997) no tags were rejected from shortnose 

sturgeon when they were coated in biologically inert material but when uncoated tags 

were used, they were rejected 33% of the time. Of those rejected, sonic tags were 

expelled within two weeks, while the radio tags were rejected within 14 weeks (Kynard 

and Kieffer 1997). Collins et al. (2002) recorded no mortality using completely internal 

tags during a three month study on tagging methods. Due to slower recovery time at 

lower temperatures, internal tags should not be implanted when water temperatures are 

below 8°C (Moser et al. 2000a, Ream et al. 2003, Kieffer and Kynard in press). Also, 

due to increased stress at higher temperatures, incisions should not be made in sturgeon 

when water temperatures exceed 27°C (Moser et al. 2000a, Kieffer and Kynard in press). 

Some researchers have experimented with an internal tag having external trailing 

antennae threaded through a permanent hole in the lateral wall of sturgeon. These tags, 

allowing for better transmission of radio frequencies, are known as Internal/External tags 

(I/E tag). However, depending on the surgical procedure used to anchor the trailing 

antennae at the exit point, certain harmful effects resulted from the chaffing and cutting 

of the trailing antenna. In one lake sturgeon I/E tagging study, Peterson and Bezold 

(2008) tagged both wild and hatchery raised fish, allowing them to recover for 14 to 21 

days prior to release. In this study, wild fish experienced 9% mortality but hatcheryreared sturgeon experienced 90% mortality. In an I/E tagging study by Collins et al.

(2002), laboratory sturgeon tagged in this manner endured large exit wounds resulting 

from the trailing antenna and eventually suffered 100% mortality. In the same study, 

internal telemetry tagging techniques and two methods of external tagging resulted in 

only one mortality. 

More recent results documented by Kieffer and Kynard (in press) found trailing antennae 

did not appear deleterious to the health of shortnose sturgeon when designed to exit the 

body wall through holes drilled in lateral scutes. Five wild fish tagged in the Connecticut 

River with I/E tags exiting through the scute were tracked for a year. All fish were 

recaptured, but the exit holes in all scutes had become larger. Until these techniques are 

better documented, NMFS recommends I/E tagging should not be done on green, Gulf, 

shortnose, or Atlantic sturgeons. 

Historically, external tags were easily shed. Collins et al. (2002) showed hatchery 

shortnose sturgeon were able to shed 100% of their external transmitters (9 cm long, 1.7 

cm diameter) when attached with a wire through the dorsal fin. However, the same 

researcher reported no external transmitter tags lost when attached to a dart tag using heat 

shrunk plastic wrap. Counihan and Frost (1999) found no external tags were shed by 

juvenile white sturgeon after one to three weeks. Sutton and Benson (2003) reported a 

14.4% shedding rate for external tags (2.1 – 4.0 cm), with 27% of the larger tags (3.4 - 

4.0 cm) shed. 
30 

More recently, researchers have documented higher retention rates with the advent of 

newer, smaller external tags and better methods of attachment (Figure 3). These external 

tags range in size between 18 and 46 mm long and only 7 to 9 mm in diameter. Using 70 

to 100 lb test monofilament line, Mike Randall and Ken Sulak (USGS, pers. comm.) 

described a method for attaching such tags bound externally to the dorsal fin using 

lightweight heat shrink electrical splice tubing and five minute, two-part epoxy. These 

researchers documented over 96% retention rates on Gulf sturgeon during 2005 to 2008 

using the following method. Their method (Mike Randall and Ken Sulak, USGS, pers. 

comm.) is described as: 

About 25 cm of monofilament is centered in approximately 20 mm length of heat 

shrink. A small quantity of epoxy is added to the tag which is then seated into the 

heat shrink tubing. The tubing is then shrunk with a heat gun until snug. This also 

warms up the mono line enough to make right angle bends at the ends of the heat 

shrink tubing. A small amount of epoxy should extrude from each end of the heat 

shrink tubing making a smooth union. Once the attachment is cooled and the epoxy 

hardened, the tag should be re-checked and the tag’s magnet affixed to the tag. A 

tape label with the identifying tag number is also wrapped around the monofilament. 

A hole is then made through the base of the sturgeon’s dorsal fin with a PIT tag 

needle which is also used as a guide to thread the mono line through the dorsal fin. 

Similarly another hole is made through the dorsal fin anterior to the first hole and the 

aft monofilament line is passed through. As the transmitter tag is pulled snugly to fit 

within the crease at the base of the dorsal fin and the body, the two monofilaments 

ends are joined on the opposite side of the dorsal fin by a short length of steel leader. 

The external tag is then secured by threading the monofilament through crimps prefastened on the ends of the steel leader. As the monofilament lines are pulled with 

opposite pressure, the leader line crimps are compressed. Finally any trailing ends of 

the monofilament or leader are cut. The leader will eventually corrode freeing the 

external tag from the fish. 

Figure 3: Location of external telemetry tag (USGS 

Southeast Ecological Science Center). 
31 

NMFS recommends acoustic telemetry tags for tracking the movements of Gulf, green, 

Atlantic, and shortnose sturgeon. NMFS would suggest tagging sturgeon externally, 

though both methods are acceptable. 

External Identifier Tags 

NMFS has authorized a variety of external-identifier tag designs and placement sites on 

shortnose sturgeon over the past 10 years. Some examples of external-identifier tags are: 

Carlin (Peterson) tags, coded wire tags, dart tags, disk anchors, double barb tag, 

elastomer, and Floy T-bar tags. Minimal research has been conducted on the effects of 

these types of tags on sturgeon species. 

The need for researchers to identify sturgeon with external-identifier tags has been called 

into question by Bergman et al. (1992) as sturgeon can be uniquely recognized by PIT 

tags. Additionally, the effectiveness and retention of these external-identifier tags is 

uncertain (Bergman et al. 1992). However, using external identifier tags can be helpful 

for identifying wide ranging sturgeon, like the Gulf, Atlantic, and green sturgeons that 

can be captured in distant locations by other researchers or commercial fishermen. 

Shortnose sturgeon are less likely to travel great distances through the ocean and into 

different rivers; therefore, external identifier tags are not as beneficial for them. 

Consequently, NMFS recommends the use of external tags to assist with the 

identification of migratory sturgeon when that information will contribute to the species’ 

recovery. 

Smith et al. (1990) compared the effectiveness of dart tags with nylon T-bars, anchor 

tags, and Carlin tags in shortnose and Atlantic sturgeon. Carlin tags applied to scutes had 

low retention rates as did dart tags; however, they also noted the dart tags caused some 

tissue damage. Carlin tags applied at the dorsal fin and anchor tags inserted in the 

abdomen showed the best retention. Although anchor tags resulted in lesions and 

eventual breakdown of the body wall if fish entered brackish water prior to their wounds 

healing, Collins et al. (1994) found no significant difference in healing rates between fish 

tagged in freshwater or brackish water. Clugston (1996) also looked at T-bar anchor tags 

placed at the base of the pectoral fins, finding beyond two years, retention rates were 

about 60%. Collins et al. (1994) compared T-bar tags inserted near the dorsal fin, Tanchor tags implanted abdominally, dart tags attached near the dorsal fin, and disk anchor 

tags implanted abdominally. He found that T-anchor tags were most effective long-term 

(92%), but also noted that all of the insertion points healed slowly or not at all and, in 

many cases, lesions developed. Collins et al. (1994) also inserted coded wire tags into 

the sturgeons’ snout and found a 100% retention after 62 days, but only 74% after two 

years, though the tags may not have been inserted deeply enough. Bordner et al. (1990) 

inserted coded wire tags deeply into the snouts of white sturgeon and found 100% 

retention after 180 days; and Isely and Fontenot (2000) also found that coded wire tags 

inserted near the dorsal fin have a 98% retention rate after 120 days. 

Winter (1983) suggested the appropriate tag weight to body weight ratio for fish was 2% 

for the tag weight in air and 1.25% for the tag weight in water. Generally, heavier tags 
32 

reduce growth or affect the swimming ability of tagged fish. But, as noted by Brown et 

al. (1999), different species of fish are better able to respond to tag weight, handling 

higher ratios of tag weight to body weight. In a tag to body weight ratio study conducted 

on lake sturgeon, Sutton and Benson (2003) recommended tag weight in air not to exceed 

1.25% of body weight. In a separate study by Counihan and Frost (1999), using the ratio 

of wet tag weight to sturgeon weight of less than 1.25%, they found the swimming 

performance of white sturgeon was affected. However, this effect was more attributed to 

the tag placement rather than the weight itself as external tags attached to the rear dorsal 

fin resulted in increased drag and unbalanced weight. Currently, NMFS is sponsoring 

directed research on a variety of sturgeon species to determine the appropriate tag to body 

weight ratio. However, until resolved, NMFS recommends not exceeding a tag to body 

weight ratio of 1.25% in water and 2% weight in air for all tags cumulatively. 

Recommendations 

General 

• PIT tags are strongly recommended to be inserted in all Gulf, green, shortnose, 

and Atlantic sturgeon without a PIT tag. 

• NMFS recommends not exceeding a tag to body weight ratio of 1.25% in water 

and 2% weight in air. 

Telemetry Tags 

• NMFS recommends I/E tagging should not be done on green, Gulf, shortnose, or 

Atlantic sturgeons. 

• NMFS recommends acoustic telemetry tags for tracking the movements of Gulf, 

green, Atlantic, and shortnose sturgeon. NMFS would suggest tagging sturgeon 

externally, though both methods are acceptable. 

External Identifier Tags 

• When appropriate, NMFS recommends the use of external tags to assist with the 

identification of migratory sturgeon. 
33 

Gastric Lavage 

The pulsed gastric lavage technique, demonstrated by Foster (1977) to sample diets of 

pickerel and largemouth bass, has not worked well for sturgeon species. This is largely 

due to the difficulty in navigating the lavage tube past the U-shaped bend of the 

alimentary canal in sturgeon, which begins after the pneumatic duct of the swim bladder 

joins the anterior end of the stomach (Figure 4, also see Haley 1998 and Brosse et al.

2002). Serious injury and mortality has occurred when lavaging sturgeon. Sprague et al.

(1993), showed gastric lavage tubes positioned prior to the pneumatic duct filled and 

burst the swim bladder and when passed beyond the bend of the alimentary canal, those 

tubes were capable of puncturing the canal and stomach lining of an unrelaxed gut. 

Haley (1998) modified the Foster (1977) protocols for gastric lavage to create a lavage 

technique appropriately safe and effective for use on sturgeon (Figure 4). Haley’s (1998) 

technique has been modified a few times with different methods created for water 

delivery into the stomach through intramedic tubing. Murie and Parkyn (2000), Savoy 

and Benway (2004), and Collins et al. (2008) each used slight variations of the water 

delivery system, but essentially used the procedures described in Haley (1998) to safely 

lavage sturgeon. NMFS recommends researchers follow these methods, as described 

below, when conducting gastric lavage of Gulf, green Atlantic, or shortnose sturgeon. 

Figure 4: Depiction of the gastric lavage technique used by Haley et al. (1998). 

First the sturgeon is anesthetized to the appropriate stage (Table 5, Stage I) causing 

the sturgeon’s esophageal and gastric muscles to relax. The sturgeon is then placed 

ventrally head down on a stretcher or sling with an irrigation tube in its mouth to 

irrigate the gills during the procedure to ensure respiration. With water running over 

the gills, a fine mesh strainer is positioned under the sturgeon’s mouth to capture the 

regurgitated contents of the stomach as it is lavaged. With the sturgeon correctly 

positioned, a soft, flexible intramedic tubing (typically polyethylene) is inserted into 

the mouth of the sturgeon and carefully directed down the alimentary canal past the 

pneumatic duct into the stomach region. At the point of resistance reached at the Ushaped bend of the stomach, the flexible tube is twisted ventrally and gently pushed 

further down the alimentary canal until the tube can be felt on the ventral surface of 

the fish. If the researcher is more conservative, the lavage procedure can begin once 

the tube reaches the point of resistance at the U-shaped bend in the stomach, as this 

method has been shown to be equally effective.
34 

Once the tube is correctly positioned, the stomach contents are evacuated with 

injected pulses of water. Haley used a syringe to inject water into the stomach, 

flushing the contents into a strainer. Variations of Haley’s technique have been used 

by other researchers to inject water using a garden sprayer holding a larger reservoir 

of water to administer the flushing, either timed (Savoy and Benway 2004) or 

manually (Collins et al. 2008). The contents are collected into an appropriately small 

meshed sieve, preserved in an alcohol filled container and the contents later identified 

in the laboratory. 

In order to conduct gastric lavage procedures, researchers should have the following 

items: 

• Garden sprayer or another appropriately sized water delivery device 

• Intramedic tubing 

• Means of anesthetization 

• 500 micrometer sieve 

• A sling or stretcher for holding the fish in the head down position 

• Jars filled with alcohol for preserving gut content samples 

Kamler and Pope (2001) and Shuman and Peters (2007) report Haley’s (1998) protocols 

are more effective for smaller fish because the syringe can only deliver a small volume of 

water. Brosse et al. (2002), Nilo et al. (2006), Savoy (2007), and Collins et al. (2008) 

developed their methods to deliver larger volumes of water to effectively lavage larger 

Atlantic and shortnose sturgeon. These researchers used varying diameter tubes and 

depending on the size of the fish, flushing slightly less than a gallon of water into the 

sturgeon’s stomach to completely evacuate its contents. 

When gastric lavage was first used with sturgeon, there were serious perceived risks to 

the individual fish. Sprague et al. (1993) reported 33% mortality (4 of 12) of white 

sturgeon they attempted to lavage. Farr et al. (2001) practiced their technique on three 

dead green sturgeon but were unable to maneuver the tubing around the bend of the 

alimentary canal. In both methods, the swim bladder filled with water resulting in 

damage to the alimentary canal and stomach. Both of these studies however used a less 

flexible aquarium tubing, a factor which potentially prevented the tubing from bending 

with the stomach and reaching the ventral portion of the stomach near the pyloric caeca. 

To avoid adverse affects in future research, NMFS recommends practicing on non-listed 

or hatchery-reared sturgeon before attempting the procedure in the wild. 

Several sturgeon researchers have also expressed concerns that delayed mortality and 

other risks associated with gastric lavage remains unknown and may not be worth the 

risks of data collection. The only way to adequately measure adverse affects is 

conducting gastric lavage on sturgeon in a laboratory setting and subsequently 

monitoring post-lavage survival, growth, and behavior. Brosse et al. (2002), Wanner 

(2006), and Mark Collins (South Carolina Department of Natural Resources, pers. 

comm.) practiced gastric lavage on captive fish with no delayed mortality prior to 

conducting lavage in the field. And in Collins et al. (2008), three Atlantic sturgeon were 

sacrificed to monitor adverse effects from lavage on wild fish. No adverse effects were 
35 

discovered. Brosse et al. (2002) reported all lavaged sturgeon were in poorer condition 

than control fish after 60 days due to weight loss. However, Collins et al. (2008) 

recaptured fish (over 70 days apart) and documented normal weight gains in the intervals 

between capture and re-lavage. Other researchers have reported successful gastric lavage 

work in the field with no immediate mortalities (Haley 1998, Brosse et al. 2002, Savoy 

and Benway 2004, Nilo et al. 2006, Guilbard et al. 2007, Nellis et al. 2007, Savoy 2007, 

Collins et al. 2008). Even if mortality is prevented by using appropriate lavage 

techniques on sturgeon, NMFS recognizes the potential risks to individual sturgeon from 

anesthesia, improper lavage technique, and individual sturgeon reacting negatively to the 

procedure. 

Recommendations

• NMFS recommends researchers follow the methods presented in this document 

and Haley (1998) when conducting gastric lavage on Gulf, green, Atlantic, or 

shortnose sturgeon. Other documents detail acceptable ways to deliver larger 

volumes of water for adult Atlantic, Gulf, and green sturgeon. 

• NMFS recommends using soft, flexible tubing (polyethylene tubing such as is 

used in hospitals) to maneuver the bend in the alimentary canal during gastric 

lavage procedures. 

• NMFS recommends practicing on non-listed or hatchery-reared sturgeon before 

attempting the procedure in the wild. 

• Sturgeon must be anesthetized to ensure relaxation of the gut walls to properly 

position gastric tubes during the procedure. 
36 

Sex Identification 

The validation of techniques to accurately identify the sex and stage of maturation of 

sturgeon that leads to the conservation of Gulf, green, Atlantic, and shortnose sturgeon 

should be a priority. All sturgeon biologists should use safe and effective methods of 

sexual identification and maturity with the fewest adverse effects to the fish’s health. 

Ideally, the sex of a sturgeon could be identified externally. A study by Vecsei et al.

(2003) examined the urogenital openings of a variety of species of male and female 

sturgeon and was able to determine the sex correctly 82% of the time. However to date, 

the sample size is too small to be confident in the methods described. 

Methods commonly used by sturgeon researchers to identify the sex and stage of 

gametogenesis of sturgeon include borescope (endoscope in the gonoduct), laparoscopy 

(endoscope through an incision in the ventral body wall), surgery and gonadal biopsy, 

ultrasound, and blood plasma. These techniques collect different information and have 

different success rates posing different risks to sturgeon. 

The safest forms of sexual identification are methods not requiring anesthetic, like 

ultrasound (Moghim et al. 2002, Colombo et al. 2004, Wildhaber et al. 2006), 

borescoping (Kynard and Kieffer 2002), plasma lipophosphoprotein analysis (Craik and 

Harvey 1984), and plasma vitellogenin analysis (Wildhaber et al. 2006). However these 

methods are time or labor intensive, as with blood and plasma analyses where researchers 

may not receive the results of their analysis until weeks later. 

Endoscopy 

Borescope 

Borescopic examination has proven an effective method for sexing sturgeon using fiber 

optic technology. Kynard and Kieffer (2002), Wildhaber and Bryan (2006), and 

Wildhaber et al. (2006) described the technique using a flexible borescope on shortnose, 

pallid, and shovelnose sturgeon where the head and body of the fish is examined under a 

lightly anesthetized condition. This procedure, lasting one to two minutes, is conducted 

with a flexible fiber optic endoscope (16cm long x 4mm diameter) inserted carefully 

through the urogenital opening and into place within the urogenital canal (Kynard and 

Kieffer 2002). Sampled females are verified by positively identifying eggs through the 

urogenital wall. Developed eggs are staged as either “early stage” or “late stage” 

individuals to identify potential spawners for the coming spring. This is done by 

carefully comparing the coloration and separation of oocytes viewed through the 

urogenital wall. Undeveloped eggs are often almond or cream-colored and sometimes 

indistinguishable from male testes, while mature eggs appear darker, separated, and well 

formed. It is noted that there are variations of this technique using a trocar to first pierce 

the genital canal to view and/or biopsy the gonads with an inserted fiber optic borescope; 

however, NMFS does not recommend this procedure on listed sturgeon. 

The above borescope is easily passed through the urogenital opening (average 7.6mm) of 

adult shortnose, juvenile Atlantic, and other sturgeon species, although there are no 
37 

similar morphological data for green sturgeon reported. Van Eenennaam et al. (2008) 

have suggested that the diameter of the urogenital canal of green sturgeon is smaller than 

other sturgeon species. The greatest potential for injury with this procedure, according to 

Kynard and Kieffer (2002), is internally at the juncture of the oviduct and urogenital 

canal, located approximately 9 to 20% of a sturgeon’s body length from the vent, 

regardless of species. The borescope must be maneuvered carefully beyond the oviduct 

to clearly see and stage eggs. However, when using a 16-cm long borescope, the probe 

tip will reach beyond the oviduct in most sturgeon of one meter length or less. Further, 

Kynard and Kieffer (2002) reported repeated probing of the oviduct valve by 4-mm and 

smaller diameter probes did not penetrate the oviduct valve or damage the urogenital 

canal regardless of species or fish length. They concluded that careful use of a properly 

sized borescope would not harm reproductive structures and would be suitable for most 

sturgeon species. 

Kynard and Kieffer (2002) examined 443 adults using a boroscope over six years. Of 

those viewed, 173 were identified as female and 270 were unidentified — either as 

females with immature eggs or identified as males. However, Wildhaber et al. (2006) 

was able to correctly identify 85% (93% accurate for males, 63% for females) of 

shovelnose and pallid sturgeon examined using a similar borescope. During their work, 

Wildhaber and Bryan (2006) and Wildhaber et al. (2006) did not document any injuries 

or mortalities associated with their borescope activities. 

Borescopy requires less time than more invasive surgery, making it a safer alternative to 

laparoscopy (described below) for field use when handling large numbers of sturgeon 

under adverse conditions. However, the borescope has limited ability to distinguish 

between females with immature eggs and male fish as compared to laparoscopy or 

biopsy. 

Laparoscope 

Several sturgeon researchers have described using laparoscopic procedures in the lab and 

field to identify the sex and egg maturity of individual sturgeon. The method for 

laboratory laparoscopy is described thoroughly by Mohler (2003), Hernandez-Divers et 

al. (2004), and Matsche and Bakal (2008). As with borescopy, the sturgeon should be 

anesthetized and held in water as much as possible. An incision (approximately 4 mm) is 

made on the ventral (Hernandez-Divers et al. 2004, Wildhaber et al. 2006) side or 

between the lateral scutes (Conte et al. 1988) of the sturgeon and the endoscope is 

inserted through the incision and maneuvered internally to allow the researcher to view 

the gonads. In Hernandez-Divers et al. (2004), the body cavities were insufflated and the 

swim bladders collapsed, but NMFS recommends avoiding either of these procedures 

when conducting laparoscopy on Gulf, shortnose, Atlantics, or green sturgeon. Although 

NMFS considers laparoscopy a more invasive endoscopic procedure than boroscopy, it is 

a more reliable method for determining the sex and stage of maturity of sturgeons 

(Wildhaber et al. 2006) and therefore recommends laparoscopy as the endoscopic 

procedure of choice. 
38 

Hernandez-Divers et al. (2004) laparoscoped 17 Gulf sturgeon. During these procedures, 

seven fish were positively identified by endoscopy alone and the other 10 were identified 

by biopsy samples of the gonad tissue. Wildhaber and Bryan (2006) examining 34 pallid 

sturgeon with both ultrasound and endoscope, positively identified the sex of 100% of the 

fish. Wildhaber et al. (2006) found that laparoscopy could positively identify the sex of 

shovelnose sturgeon 93% of the time (93% for males, 92% for females). 

Adverse effects were not reported in any of the papers discussing laparoscopy. 

Hernandez-Divers et al. (2004) reported 100% survival after extensive surgeries (45 

minutes to an hour) for their 17 Gulf sturgeon. Unfortunately this work was conducted in 

a controlled laboratory setting by three surgeons and does not represent typical field 

research conditions. Additional research determining adverse effects associated with 

laparoscopic procedure still need to be documented, particularly on gravid females 

captured prior to initiating a spawning run. Several researchers have reported capturing 

sturgeon can may be related to abandoned spawning runs (Moser and Ross 1995, Kynard 

et al. 2007), but there have been no studies addressing the effects of anesthesia or 

laparoscopy on mature, late stage females still occupying their winter staging habitat 

prior to spawning. 

Surgical Biopsy 

Surgical biopsy and histological examination of a sturgeon’s gonadal tissue is the most 

accurate while also the most invasive way to identify the sex and stage of maturity of a 

sturgeon (Van Eenennaam et al. 1996, Van Eenennaam and Doroshov 1998, Fox et al.

2000, Webb and Erickson 2007, Flynn and Benfey 2007). Chapman and Park (2005) 

conducted gonad biopsies on Gulf sturgeon by anesthetizing them and placing them in a 

sling on their backs. A two to four cm ventral incision was made, after which, a small 

gonadal biopsy was removed (Chapman and Park 2005, Webb and Erickson 2007). 

Surgical biopsy, usually removing about 1 cm3

 of tissue (Fox et al. 2000, Webb and 

Erickson 2007), lasts two to three minutes (Chapman and Park 2005). After biopsies are 

completed, the gonadal tissue is microscopically examined to verify the sex as well as the 

precise stage of maturation of sturgeon (Van Eenennaam et al. 1996, Van Eenennaam 

and Doroshov 1998). 

As with other forms of surgery, the risks are minimized when performed in the laboratory 

but there is little to no information available on the extent of infection or delayed 

mortality. Although there is documentation of surgically sterilized sturgeon regenerating 

gonadal tissue, there is little information regarding the loss of reproductive potential due 

to the removal of small samples of gonadal tissue (Kersten et al. 2001, Hernandez-Divers 

et al. 2004). And, while it is known that the gonads deliver hormones to the fish that 

influence behavior (Hernandez-Divers et al. 2004), there have been no studies dealing 

with potential changes in behavior from small losses of gonadal tissue. Chapman and 

Park (2005) monitored Gulf sturgeon for 30 days following biopsy and reported no 

mortality. 

In situations when knowing the stage of gametogenesis could lead to recovery of the 

listed species, laparoscopy or biopsy would be appropriate, but due to the increased risk 
39 

of these procedures, NMFS only recommends using these procedures in a laboratory 

setting. If there are situations when these methods would be more likely to contribute to 

the recovery of these species than other available methods, NMFS would recommend 

their use under limited circumstances. Gonadal biopsy should only be performed in the 

field opportunistically while a researcher is implanting an acoustic tag. 

Ultrasound 

One of the safest and least invasive methods of sexual identification is the use of 

ultrasound. These devices, although costly, allow researchers to observe the sex organs 

of sturgeon without surgical incision or sedation. Ultrasound is the technique with the 

most potential and is becoming more accurate as both technologies improve and readers 

become more experienced (Joel Van Eenennaam, University of California Davis, pers. 

comm.). 

When conducting ultrasound analyses, the procedures described by Wildhaber et al.

(2006), or slight variation of these techniques, appear to be the safest described in the 

literature. Sturgeon are placed in a prone position in a tank of water with their ventral 

surfaces exposed to air. The ultrasound transducer is coated with ultrasound gel and then 

covered in a protective plastic sheath to prevent any scratches to the ultrasound from the 

sturgeon’s scutes. During scanning, output power, focus depth, and frame rate are kept 

constant. The transducer is maneuvered along the abdomen between the gills and the 

anus, keeping the wide end of the transducer facing the head and tail. The ultrasound 

cannot penetrate the hard calcium of the scutes, so there is no reason to attempt to 

ultrasound the sides or back of the sturgeon (Wildhaber et al. 2006). 

Moghim et al. (2002) examined 249 anesthetized stellate sturgeon with ultrasound and 

then performed necropsies to verify the accuracy of the ultrasound. Overall, ultrasound 

was 97.2% accurate in determining sex with the procedure taking only 30 seconds to 

complete. Mature females were the easiest to identify (100%), followed by immature 

females (99.3%), mature males (96.5%), and then immature males (76.2%). Colombo et 

al. (2004) examined 51 euthanized shovelnose sturgeon and determined ultrasound was a 

viable method of sex identification. They were able to correctly identify the sex of 

sturgeon 88% of the time, though only 40% of post-spawned females were accurately 

identified. Excluding post-spawned female sturgeon, the ultrasound correctly identified 

the sex of sturgeon 94% of the time. Additionally, Wildhaber and Bryan (2006) 

accurately identified the sex of 100% of pallid and shovelnose sturgeon using ultrasound 

coupled with borescope. In another study, Wildhaber et al. (2006) correctly identified 

only 68% of fish in the field and 70% of fish in the laboratory. In both of these cases, 

males were more often correctly identified, which is similar to the results from Colombo 

et al. (2004) but opposite the findings from Moghim et al. (2002). 

When performed without anesthesia, there are no risks associated with ultrasound 

examination of sturgeon. However, while ultrasound is able to identify gender, it is not a 

promising method for determining the stage of eggs. When working with listed Gulf, 

shortnose, Atlantic, and green sturgeon, NMFS generally recommends using ultrasound 

for instant sexual identification of fish in the field. This method is the least stressful and 
40 

comparably accurate to other available methods that provide immediate identification. 

Due to the expense of ultrasound technology, boroscoping shortnose, Gulf, and Atlantic 

sturgeon is an acceptable alternative. More research is needed to determine if 

boroscoping is safe for green sturgeon. 

Blood Plasma 

Potentially one of the most promising, most accurate, and least stressful procedures used 

to sex sturgeon is an analysis of blood plasma. Researchers have used vitellogenin or sex 

steroids such as testosterone, 11-ketotestosterone, and estradiol to assess the sex and 

stage of maturity for pallid, shovelnose, hybrid bester, and white sturgeon (Amiri et al.

1996, Webb et al. 2002, Wildhaber et al. 2006). 

Blood samples are obtained from the caudal vein (Figure 5) and centrifuged to isolate the 

plasma where it is then analyzed by radioimmunoassay or frozen for later analysis. In 

initial studies, testosterone was used to discern sexual maturation (79% accuracy for 

males, 85% for females), as it is significantly elevated in mature male and female 

sturgeon (Webb et al. 2002). If testosterone indicates the sturgeon is maturing, estradiol 

levels of female white sturgeon exceed 2 ng/ml 93% of the time, while males and 

immature white sturgeon estradiol levels never exceed 2 ng/ml (Webb et al. 2002), 

resulting in reasonably accurate identification of immature males (72%), immature 

females (88%), mature males (96%), and mature females (98%). Later, researchers 

studied vitellogenin along with the sex steroids testosterone and estradiol (Wildhaber et 

al. 2006). At all stages of development, vitellogenin was significantly elevated in 

females when compared to males, predicting the sex of the sturgeon with over 99% 

accuracy. After sex determination, the same steps taken by Webb et al. (2002) can 

determine whether each gender of fish is sexually mature, as estradiol is significantly 

higher in maturing females and ketotestosterone is significantly higher in maturing males. 

Figure 5: Blood collection from a shortnose sturgeon. 

Photograph by J. Gibbons, SCDNR 

Techniques for blood plasma analysis show promise in identifying sex and egg 

maturation of sturgeon, and should continue to be evaluated for use on Gulf, shortnose, 

Atlantic, and green sturgeon. However, this technique can only identify the sex and stage 

of maturity of a sturgeon after the sturgeon has been captured and released. Therefore 
41 

this technique is not useful if researchers only need to know the sex of a sturgeon to 

identify optimal fish for an acoustic tag. If the sex of the fish is not needed immediately, 

but rather for later population analyses, blood samples are the preferred method. 

Ultrasound would also be an acceptable method even if the results are not needed 

immediately. These methods are least stressful and highly accurate in this situation. 

Recommendations 

Endoscopy 

• During borescope procedures, NMFS does not recommend using a trocar to first 

pierce the genital canal to view and/or biopsy the gonads. 

• Althought NMFS considers laparoscopy a more invasive endoscopic procedure 

than boroscopy, it is a more reliable method for determining the sex and stage of 

maturity of sturgeons (Wildhaber et al. 2006) and therefore recommends 

laparoscopy as the endoscopic procedure of choice. 

Gonadal Biopsy 

• NMFS does not recommend the use of laparoscopy or biopsy on wild Gulf, green, 

Atlantic, or shortnose sturgeon, but does recommend their use on hatchery and 

laboratory sturgeon. However, if there are situations when these methods would 

be more likely to contribute to the recovery of these species than other available 

methods, NMFS would recommend their use under limited circumstances. 

• Gonadal biopsy should only be performed in the field opportunistically while a 

researcher is implanting an acoustic tag. 

Ultrasound 

• NMFS generally recommends using ultrasound for instant sexual identification of 

fish in the field. 

Blood Plasma 

• Blood samples are the preferred method for determining the sex and stage of 

maturity of sturgeon when that information is not needed at the time of sampling. 
42 

Age Estimation 

Age estimates of sturgeon populations help researchers and managers understand 

sturgeon growth rates, ages at maturity, mortality rates, productivity, longevity, and year 

class strength (Campana 2001, Paragamian and Beamesderfer 2003). Such knowledge is 

critical for designing appropriate fisheries management policies. 

Bony structures form opaque and transparent age rings each year in most fish species in 

response to changes in temperature or other annual cycles. These rings, or annuli, are 

roughly correlated to sturgeon age. Unfortunately, most bony structures, such as 

clavicles, cleithra, opercles, and medial nuchals are not options for listed species of 

sturgeon because such sampling is lethal (Brennan and Cailliet 1989, Stevenson and 

Secor 1999, Jackson et al. 2007). Other structures such as dorsal scutes and pectoral fin 

spines, so named because of a dermal bone sheath (Feindeis 1997), are more viable 

options, but scutes are more difficult to read than fin spines (Huff 1975, Brennan and 

Cailliet 1989, Stevenson and Secor 1999, Jackson et al. 2007). 

Pectoral fin spines are sampled by researchers similarly across the United States. The 

following methodology is therefore recommended for sampling pectoral fin spines of 

Gulf, green, Atlantic, and shortnose sturgeon (Figure 6). 

Using a hacksaw or bonesaw, two parallel cuts are made across the leading pectoral 

fin spine approximately 1-cm deep. The blade of the first cut is positioned no closer 

than 0.5-cm from the point of articulation of the flexible pectoral base to avoid an 

artery at this location (Rien and Beamesderfer 1994, Rossiter et al. 1995, Collins and 

Smith 1996). The second cut is made approximately 1-cm distally (Everett et al.

2003, Fleming et al. 2003b, Hurley et al. 2004, Hughes et al. 2005), where a pair of 

pliers can be used to remove the resulting fin spine section. The section is then 

placed in an envelope and air-dried for several days or weeks. Later it is cut into thin 

slices (usually about 0.5 to 2 mm thickness) typically using a jeweler’s saw or a 

double bladed saw (Stevenson and Secor 1999, Everett et al. 2003, Fleming et al.

2003b, Hurley et al. 2004, Hughes et al. 2005, Johnson et al. 2005, Collins et al.

2008). The sections are then mounted onto the substrate of choice including clear 

glue, fingernail polish, cytosel, or thermoplastic cement. The cross-section detail of 

the fin spine annuli are then studied using stereoscopic readers. 
43 

Figure 6: Diagram of the appropriate method for removing a small section of fin spine for 

age analysis.

Accuracy and Precision of Estimates 

Accuracy and precision of the fin spine age estimates are concerns of fishery biologists 

and management agencies. Precision is a measurement of the distance between two 

reader’s interpretations of the same fin spine sample, while accuracy is a measurement 

between the reader estimate of a sturgeon’s age and the actual age (Beamish and 

MacFarlane 1983, Campana et al. 1995, Campana 2001, Hurley et al. 2004). To estimate 

precision, mark-recapture studies, oxytetracycline chemical marking studies, hatchery 

release studies, and in hatchery studies have been conducted to validate the age 

estimation process and also verify the assumption of one opaque and one translucent ring 

are formed each year (Clugston et al. 1990, Rien and Beamesderfer 1994, Campana et al.

1995, Rossiter et al. 1995, Stevenson and Secor 1999, Campana 2001, Paragamian and 

Beamesderfer 2003, LeBreton and Beamish 2004, Hurley et al. 2004, Jackson et al.

2007). Most studies of age estimates measure precision using at least two individual 

readings of the same slide. Subsequently, either the variability is recorded between 

readers, or the differences in reader’s estimates are reconciled immediately after 

measurement. 

Most age estimation studies suggest the results obtained should be used with caution 

because, while fin spines may provide the safest and most accurate estimation of age, 

they also consistently underestimate the actual age. The typical sources of error reported 

have been: 1) the rings are too close together or not clearly differentiated; 2) the original 

ring is difficult to identify; 3) the rings are missing within deteriorating sections; or 4) 

secondary fin spines, split rings, false rings, or spawning bands tend to create more or 

fewer rings than the actual age (Nakamoto 1995, Rossiter et al. 1995, Lai et al. 1996, 

Stevenson and Secor 1999, Farr et al. 2001, LeBreton and Beamish 2004, Whiteman et 

al. 2004). Moreover, fin spines from hatchery fish are often shaped differently, resulting 

in a more difficult age comparison control. 

Accuracy of Estimates

The accuracy of fin spine estimates has been measured for Atlantic, pallid, shovelnose, 

white, lake, and Gulf sturgeon. Rossiter et al. (1995) and Stevenson and Secor (1999) 

monitored fish after one to three years between capture and found for lake and Atlantic 
44 

sturgeon respectively, growth rings did develop once a year. But LeBreton and Beamish 

(2000) determined only five of seven populations of lake sturgeon exhibited a series of 

one opaque and one translucent ring formed per year. This was also seen by Morrow et 

al. (1998) who documented two bands forming annually in shovelnose sturgeon fin 

spines during warmer years. Van Eenennaam et al. (1996) showed reader error of one to 

two years underestimation for Atlantic sturgeon at true ages ranging between 15 and 30 

years. For shovelnose sturgeon and Atlantic sturgeon, age estimates underestimate actual 

age. The age underestimation was 1.6 years for fish under 15 years, 1.7 years for fish 

between 16 and 20 years, and 4.3 years for fish over 21 (Whiteman et al. 2004). A 

similar result was reported by Paragamian and Beamesderfer (2003) and also by Rien and 

Beamesderfer (1994) for white sturgeon, each finding age underestimation for white 

sturgeon under 60cm was over 70%, while the accuracy fell to below 60% for fish above 

100cm. Moreover, using length to estimate age of sturgeon has proven unreliable. 

Clugston et al. (1990) recorded lengths of Gulf sturgeon after one year in the laboratory 

and then noted the inconsistent growth of fish in the wild during all months of the year. 

They concluded fish of similar sizes captured in the wild yield variable growth rates, 

suggesting length at age charts are flawed because growth is not constant among 

individuals. Paragamian and Beamesderfer (2003) provided additional evidence of 

invalid length at age charts using wild sturgeon. However, Peterson et al. (2000) and 

Schueller and Peterson (in press) demonstrated that juvenile sturgeon younger than three 

can be aged using length at age charts. 

In the most extensive mark-recapture study to date, analyzing sturgeon at large over five 

years, Paragamian and Beamesderfer (2003) examined 760 marked (known age) white 

sturgeon recaptured up to 23 years later. They found ages were underestimated between 

30 and 60%, depending on the time spent at large, meaning that age estimates were 1.5 to 

2 times below the actual age of the fish. For marked-recaptured shortnose sturgeon, there 

was 96% accuracy between the readers’ age estimates and the time the sturgeon spent at 

large. However, when using known-age fish, only 34% of the readers’ estimates were 

accurate within one year (Collins et al. 2008). Also, when using multiple slides from the 

same fin spines of known-age hatchery fish, Hurley et al. (2004) reported only 28% of 

the estimates were correct, while 56% were within one year and 89% were within two 

years. 

Precision of Estimates

As discussed previously, when measuring the precision of fin spine aging estimates, 

multiple readers estimate the age of identical sturgeon fin spines and then their results are 

compared to determine the variance between readers’ estimates. Fleming et al. (2003b) 

studied 88 shortnose sturgeon fin spines where multiple readers were able to reach an 

agreement after consultation 100% of the time. Everett et al. (2003) analyzed shovelnose 

sturgeon using multiple readers and found the readers could not reach agreement on 26 of 

736 (3.5%) of the samples when they attempted to reconcile measurements. Rossiter et 

al. (1995) also showed agreement between reader measurements while analyzing 20 lake 

sturgeon. They found high precision between readers for fish under 15 years old; 

however, for fish over 18 years old, reader agreement dropped to 80%. In the first two 

studies mentioned above, the readers reconciled measurements when there was a 
45 

disagreement in age estimation, while the latter study was conducted on only 20 samples 

without reconciliation of estimates. 

While some studies have found general precision and agreement between readers, others 

were less successful. Van Eenennaam et al. (1996) showed multiple readers agreed on 

readings of Atlantic sturgeon fin spine samples approximately 33 to 40% of the time. 

Stevenson and Secor (1999) also evaluated reader agreement of Atlantic sturgeon fin 

spines and found no significant difference, but the disagreement error was approximately 

1.2 years on average between readers. Nakamoto (1995) analyzed 154 green sturgeon fin 

spines and found readings from 34% differed by fewer than two years and 66% of the 

readings differed by fewer than five years. Rien and Beamesderfer (1994) measuring 935 

white sturgeon fin spines twice, found only 37% agreement between readers and 68% 

agreement within one year. Jackson et al. (2007) found 80% of the time multiple readers 

estimated the age of shovelnose sturgeon within one year and 100% were estimated 

within two years. However Whiteman et al. (2004) found reader agreement on 234 

shovelnose sturgeon samples of only 18% and within one year was still only 46%. 

NMFS recommends all sturgeon age estimates derived from fin spine analysis should test 

for precision between readers. 

As discussed above, one major assumption for fin spine age estimation is each fin spine 

develops a ring each year; but there is evidence to suggest each fin spine may be 

different. Jackson et al. (2007) simultaneously removed both fin spines from shovelnose 

sturgeon and showed the spines from the same fish resulted in the same estimated age 

36% of the time, within one year 66% of the time, and within two years 84% of the time. 

But this could be a result of how the fin spines are prepared, as measurements of 64 slides 

made from 16 pallid sturgeon fin spines resulted in only 25% agreement from the same 

spines (Hurley et al. 2004). Jackson et al. (2007) concluded the preparation of fin spines 

must be standardized so results can be reproducible. 

Age Validation 

Several researchers have suggested slow growth of adult and pre-spawn females may 

explain why some fin spine rings are closely spaced and become more closely spaced as 

fish get older (Beamish and MacFarlane 1983, Nakamoto 1995). It is thought the 

distance between rings is influenced by changes in food supply, metabolism, behavior, 

and environmental conditions as the sturgeon mature. 

Accordingly, sturgeon researchers have begun to develop age estimate correction factors 

to validate age estimates of populations of different species. Bruch et al. (2009), while 

researching lake sturgeon, found growth increments on pectoral fin spine cross sections 

underestimated true age of fish older than 14 years and error increased with age, whereas 

otoliths accurately estimated true age up to at least 52 years. Increment formation in 

juvenile lake sturgeon pectoral fin spines was clearer and easier to interpret than otoliths. 

A power function developed by Bruch et al. (2009) provided a means for correcting 

existing age estimates obtained from lake sturgeon pectoral fin spines. For that reason, 

NMFS recommends using salvage specimens of Gulf, green, Atlantic, and shortnose 

sturgeon to establish age estimation correction factors. 
46 

Deleterious Effects of Fin Spine Sampling 

Kohlhorst (1979) first reported potentially deleterious effects of fin spine removal from 

white sturgeon during a mark-recapture study where an incidence of mortality was 

recorded. The percentage mortality reported could have been magnified by a small 

sample size, but concern over this result triggered additional research in the laboratory. 

Collins et al. (1995) and Collins and Smith (1996) monitored the effects of fin spine 

removal of juvenile shortnose and Atlantic sturgeon in a laboratory. Removing the entire 

leading fin spine from the base, a method not currently recommended for sampling finrays, they found wounds healed rapidly and that the remaining secondary pectoral fin 

spine grew in circumference until appearing very similar to the original fin spine. There 

were no significant differences for growth or survival between treatment and control 

sturgeon. 

In other laboratory studies testing fin spine function, Wilga and Lauder (1999) found 

pectoral fins function by orienting the body vertically in the water column, but they are 

not used during locomotion. Following this study, Parsons et al. (2003) removed pectoral 

fin spines from shovelnose sturgeon placing them in tanks, where the current could then 

be increased to test their ability to hold position in a current. Without fin spines, 

treatment sturgeon were able to hold their position in a current as well as control 

sturgeon. 

Most recently, while conducting mark-recapture surveys of Atlantic and shortnose 

sturgeon, Collins et al. (2008) discovered secondary fin spines had grown abnormally on 

older, mature Atlantic sturgeon after the leading fin spine had been taken months earlier. 

Concluding this regrowth could be due to slower growth of mature, adult fish and 

possibly become detrimental to the sturgeons’ health, their team no longer samples fin 

spines from larger, adult sturgeon. Because of increased error in reading fin spines of 

older fish and evidence of abnormal regrowth, NMFS does not recommend taking fin 

spine samples from mature Gulf, shortnose, Atlantic, or green sturgeon. 

Alternative Methods for Age Estimation 

NMFS recommends developing newer, more accurate and precise methods of aging Gulf, 

green, Atlantic, and shortnose sturgeon. In recent years, Bruch et al. (2009) analyzed the 

use of radiocarbon bombing to estimate the ages of lake sturgeon using otolith cores. 

This is not a non-lethal technique, but if further testing indicates using other bony 

structures such as scutes for accurate and precise age estimates, this may become a useful 

method for age estimation. Likewise, telomeres have recently been used to estimate fish 

age. Hatakeyama et al. (2008), testing small teleost fish, found that telomere length 

shortens through the life of the fish and is inversely related to the length of the fish. 

However, no change in telomere length was noted for European sea bass between 12 and 

94 months of age (Horn et al. 2009). Specific studies should be conducted on sturgeon to 

determine if telomere analysis could determine the age of sturgeon. 
47 

Recommendations 

General 

• NMFS recommends removing a 1cm portion of the pectoral fin spine from just 

above the point of articulation to estimate the age of Gulf, green, Atlantic, and 

shortnose sturgeon. 

Accuracy and Precision of Estimates 

• NMFS recommends fin spine derived age estimates be used with caution because 

they consistently underestimate the actual age. 

• NMFS recommends all sturgeon age estimates derived from fin spine analysis 

should test for precision between readers. 

• NMFS acknowledges the preparation of fin spines must be standardized so the 

results are reproducible and encourages future research to achieve this goal. 

Age Validation 

• NMFS does not recommend using lethal methods or length/age charts to estimate 

ages of Gulf, Atlantic, green, or shortnose sturgeon, except when working with 

juvenile sturgeon under three years of age. 

• NMFS recommends using salvage specimens of Gulf, green, Atlantic, and 

shortnose sturgeon to establish age estimation correction factors. 

Deleterious Effects of Fin Spine Sampling 

• Because of increased error in reading fin spines of older fish and evidence of 

abnormal regrowth, NMFS does not recommend taking fin spine samples from 

mature Gulf, shortnose, Atlantic, or green sturgeon. 

Alternative Methods for Age Estimation 

• NMFS recommends developing newer, more accurate and precise methods of 

aging Gulf, green, Atlantic, and shortnose sturgeon. 
48 

Salvage Specimens 

Dead or salvaged specimens can be invaluable for a number of basic and applied aspects 

of sturgeon biology and conservation. Scientific uses include, but are not limited to, 

morphology, genetics, histopathology, contaminants, age and growth, food habits, 

cryopreservation of sperm, and human impact/anthropogenic mortality. Educational uses 

of sturgeon collected include, but are not limited to, taxidermy, collection of hard parts 

(e.g., scutes, bones, and entire skeleton), necropsy, and development of sampling and 

necropsy procedures and manuals. 

Although it is important to maintain salvaged specimens and their derivative tissues, 

making them available for future researchers and educators, listed sturgeon are protected 

and transfer of specimens must still be carefully documented under the ESA. 

Persons/laboratories receiving specimens must be authorized to possess listed species. 

All sturgeon research permits issued by NMFS currently include provisions for 

preserving incidental mortality resulting from research or found opportunistically. 

If dead Gulf, green, shortnose, or Atlantic sturgeon are found or a researcher has a need 

for salvaged sturgeon or sturgeon parts, contact NMFS Headquarters in Silver Spring, 

Maryland at (301) 713-2289. 
49 

Acknowledgements

We thank all of the Gulf, green, Atlantic, and shortnose sturgeon researchers who 

provided data, information, and comments throughout the development of these 

protocols. We are specifically grateful to Boyd Kynard, Micah Kieffer, Doug Peterson, 

Mark Collins, Bill Post, Tom Savoy, Gayle Zydlewski, Steve Fernandez, Joe Hightower, 

Matt Balazik, Chris Hager, Mike Mangold, David Secor, Steve Minkkinen, Hal 

Brundage, Dewayne Fox, Matt Fisher, Jerre Mohler, Mark Bain, Gail Whippelhauser, 

Jeff Wilcox, Ken Sulak, Mike Randall, Frank Parauka, Marty Gingras, Ryan Mayfield, 

Jason DuBois, Joel Van Eenennaam, Tom Rien, Dan Erickson, Mary Moser, Olaf 

Langness, Peter Allen, Mike Parsely, and Tim King. We are also grateful for the 

assistance provided by Stephania Bolden, Kelly Shotts, Russell Bohl, Lynn Lankshear, 

Kim Damon-Randall, Jessica Pruden, Melissa Neuman, Susan Wang, and David 

Woodbury. 
50 

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