Reproductive Parameters in Invasive Blue Catfish (Ictalurus furcatus) From Tributaries of the Chesapeake Bay in Maryland and Delaware, 2020–22
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Acknowledgments
Technical and logistical support for the field collection of specimens was provided by Brett Coakley, Mary Groves, Tim Groves, and Dr. Joseph Love of the Maryland Department of Natural Resources. Laboratory technical support for the preparation of histological specimens was provided by Adam Sperry, formerly of the U.S. Geological Survey. Funding support of this project was provided by the Chesapeake Bay Program and the U.S. Geological Survey Biological Threats and Invasive Species Research Program. The findings and conclusions in this report are those of the authors and do not necessarily represent the views of the U.S. Fish and Wildlife Service.
Abstract
Over the past few decades, Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish) have become a formidable invasive species in tidal tributaries of the Chesapeake Bay in Maryland and Delaware. Knowledge of their reproductive behaviors can support managers in the determination of ideal timing and implementation of mitigation strategies. In 2020–22, the U.S. Geological Survey sampled blue catfish from the Chesapeake Bay’s tidal reaches of the Nanticoke River, Broad Creek, Marshyhope Creek, and Patuxent River in Maryland and Delaware from March to October. All fish were analyzed with histology to assess reproductive stages (immature, pre-spawn [early and late], and post-spawn). Plasma was collected for multiple endpoints including 17β-estradiol (E2), calcium, and total protein. Results indicated that female spawning generally occurred from late April through June, as evidenced by the histological data showing that the number of vitellogenic oocytes in late pre-spawn females began to increase in April, peaked in May, and gradually declined through July. In males, the greatest number of late pre-spawn individuals was observed in April and gradually declined through June. Additionally, female E2 levels were highest in late, pre-spawn females, thus showing a similar trend as the histological results, indicating that this endpoint can be used for assessing reproductive changes over time. Collectively, this study documents typical spawning patterns in blue catfish within the Chesapeake Bay watershed. However, further research across different watersheds would enhance data availability and inform more comprehensive management strategies.
Introduction
Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish) are identified among federal and state partners in the mid-Atlantic region as a priority, if not the top priority, when considering the management of aquatic invasive species (Densmore, 2020). Since their intentional introduction in the James River in Virginia (fig. 1) almost 50 years ago, these fish have disseminated widely throughout lower reaches of other tributaries in Virginia and Maryland, infiltrating throughout Maryland’s tidal basin (Chesapeake Bay Program, 2020). Between 2020 and 2023, the number of rivers where they have been detected has nearly doubled (Maryland Department of Natural Resources, 2023). Blue catfish have the potential to grow to a large size (1,650 mm and 45,000 g) and live 20 or more years. They can adapt to a wide range of habitats and feed opportunistically on a varied diet, making them the apex predators in the habitats they occupy (Schlosser and others, 2011). However, the life history characteristics that have allowed the blue catfish to dramatically expand their range make them a threat to important regional fisheries such as Morone saxatilis (Walbaum, 1792; striped bass), Alosa sapidissima (Wilson, 1811; American shad), and Callinectes sapidus (Rathbun, 1896; blue crab), among others.
Information related to the reproduction of invasive blue catfish in the mid-Atlantic region would help managers in the Chesapeake Bay area better understand their spawning habits and support modeling for population dissemination risks. Although some information has been gathered pertaining to blue catfish spawning and reproductive parameters in the large tributaries in Virginia (Nepal and Fabrizio, 2021), comparative reproductive assessments across other, smaller tributaries would provide valuable evidence regarding spawning, fecundity, and the potential for applying reproductive endpoints for early detection or mitigation strategies. Conducting reproductive assessments could further inform fisheries managers on temporal and spatial variabilities in spawning throughout the region.
To achieve this goal, the U.S. Geological Survey sampled blue catfish from the Nanticoke River, and two of its tributaries, Broad Creek and Marshyhope Creek (fig. 1), in Maryland and Delaware in 2020–22, and analyzed plasma for 17β-estradiol (E2), total calcium, and total protein. These endpoints are reliable indicators of the reproductive status in fishes because of their significant increase during vitellogenesis prior to spawning. E2, a hormone secreted by the ovary, travels through the blood binding to estrogen receptors in the liver to initiate the production of vitellogenin (an egg-yolk precursor protein; Reading and others, 2017). Thus, as a proxy for direct measurement of vitellogenin, E2 and calcium (Gillespie and de Peyster, 2004) and total protein (Ogŭz and Ünal, 2012) were measured in the plasma. Additionally, blue catfish gonads were histologically processed to assess reproductive development from these sites as well as a site along the Patuxent River, Maryland, in 2020 (fig. 1).
Methods
Methods for this study were developed to analyze seasonal changes in blue catfish reproductive status. This study included the use of histology to examine the status of gonadal development and the analysis of plasma endpoints associated with reproduction. These methods are described in detail in this section.
Fish Collection
Blue catfish were collected by multiple methods, including boat electroshocking, gill nets, trot lines, and trawling by the Maryland Department of Natural Resources, Salisbury University in 2022, Delaware Department of Natural Resources and Environmental Control, and Division of Fish and Wildlife in 2020–21, and by hook and line during a fishing tournament at Marshyhope and Broad Creeks (fig. 1). Fish were sampled from the Nanticoke River (a Chesapeake Bay tributary; fig. 1) in July–October 2020, June 2021, and March–May and July–October 2022. All blue catfish sampled from the Nanticoke River were analyzed with histology and for plasma endpoints. Additional blue catfish analyzed with histology were sampled from two Nanticoke River tributaries, Marshyhope Creek in March–May and July–September 2022 and Broad Creek in June 2021, and the Patuxent River (a Chesapeake Bay tributary) in September 2020 (fig. 1).
Map showing the site locations where Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish) were sampled from tributaries of the Chesapeake Bay in Maryland and Delaware in 2020–22.
Morphometrics and Necropsy
All blue catfish were weighed to the nearest gram (g) and measured for total length millimeter (mm). Fish were euthanized by immersion in an ice bath followed by cranial concussion and pithing according to the U.S. Geological Survey (USGS) Eastern Ecological Science Center’s (EESC; #2020-FHB-001) and Salisbury University’s Institutional Animal Care and Use Committee (#SU-0065) standards. For selected (greater than 300 mm) blue catfish sampled from the Nanticoke River and Marshyhope Creek, blood was drawn from the caudal vessel with a 20–22-gauge needle with a 1–5 cubic centimeters (cc) syringe (depending on fish size), expelled into a heparinized vacutainer (BD Vacutainer, Fisher Scientific, Hampton, N.H.), and held on wet ice less than or equal to (≤) 4 hours until returned to the USGS EESC Leetown Research Laboratory, Kearneysville, W. Va. At the lab, the samples were centrifuged at 840 relative centrifugal force (rcf) for 10 minutes to separate and collect the plasma. The plasma was stored in microcentrifuge tubes at −80 degrees Celsius (°C) until analysis. For all blue catfish, a slit was cut along the ventral side from the anus to the operculum and pieces of gonad were removed and placed in 10 percent neutral buffered formalin for histological analysis.
Plasma Analyses
Plasma Preparation and Steroid Extraction
Plasma samples were shipped overnight on dry ice to the USGS’s EESC S.O. Conte Anadromous Fish Research Laboratory (Turner’s Fall, Maine) and held at −80°C until sample processing. Three separate plasma aliquots were prepared for plasma E2, and total calcium and protein to account for sample variability due to potential storage concerns caused by freeze-thaw cycles.
From thawed plasma samples, E2 was extracted by using standard ethyl ether extraction (Hwang and others, 1992). Briefly, 250 microliter (𝜇l) sample aliquots were dissolved in 10 times the volume of diethyl ether (Sigma-Aldrich, St. Louis, Mo.) and vortexed thoroughly for 30 seconds. Samples were centrifuged for two minutes at 840 rcf and frozen at −80°C for 10 minutes, and supernatants were transferred to clean, 2 milliliter (mL) tubes. This process was repeated to ensure maximum hormone extraction. Elutes from each sample were combined in one 2 mL tube and evaporated using a heat block at 40°C for 15 minutes. Samples were resuspended in distilled water and frozen at −80°C until further analysis.
Plasma E2 was measured with a commercially available ELISA assay kit (Eagle Biosciences Estradiol ELISA Assay Kit, ESD31-K01, Amherst, N.H.), following manufacturer’s protocol. The E2 ELISA kit was validated for use in blue catfish by running parallelism, accuracy, and precision tests (performing multiple runs of the same sample) using mature and immature female samples, as outlined by Metcalfe and others (2018). To ensure E2 in samples remained within the detection range, samples were diluted 1:5 volumes in ultrapure water (Molecular Biology Grade, BP2819, Fisher Scientific, Rockford, Ill.). Standards and samples were run in duplicate. The linearity of sample and standard dilutions was confirmed and compared by linear regression. Precision tests were conducted to determine intra-assay (within plate) and inter-assay (different plates) variation. Both variations were calculated as the coefficient of variation and were shown to be ≤10 percent, well below the acceptable criteria of 15 percent (Metcalfe and others, 2018).
Plasma Calcium
Plasma calcium was measured using a commercially available quantitative colorimetric calcium kit (QuantiChrom Calcium Assay Kit, DICA-500, BioAssay Systems, Hayward, Calif.). The assay was conducted in a 96-well clear, flat-bottom high bind microplate (Item 3369, Corning, Corning, N.Y.). Standards and working solutions were prepared immediately prior to use. Endpoint optical density was read at 612 nanometers. The colorimetric assay was validated in immature and mature male and female plasma samples using parallelism, accuracy, and precision tests. Serial dilutions of samples were found to be linear to standard serial dilutions. The optimal volume was 5 µL per sample loaded. The accuracy was appropriate, with percent spiked recovery from kit standard equal to 105 percent and from previously run samples equal to 101±2 percent in males and 112±3 percent in females. Precision within plates and across plates had a coefficient of variation of ≤10 percent.
Plasma Protein
Plasma protein concentration was determined with a quantitative colorimetric bicinchoninic acid (BCA) protein assay (Pierce, Fisher Scientific, Rockford, Ill.) according to manufacturer’s protocol. Using a 96-well clear round-bottom microplate (Item 3797, Corning, Corning, N.Y.), 10 µL aliquots of standards and samples were incubated with BCA reagents for 30 minutes at 37°C. The endpoint of optical density was read at 562 nm on the previously mentioned microplate reader. The assay was validated for use in immature and mature males and females. Serial dilutions of plasma were linear to kit BSA standard curve following a 1:50 dilution. The percentage of spiked recovery of the BCA protein assay was 92 percent in the kit standard, 98±3 percent in males, and 117±3 percent in females. Thus, these values were well within the acceptable range of 80–120 percent recovery. The coefficient of variation was ≤10 percent at this dilution for all standards and samples across and within plates.
Histology
Pieces of gonad (taken from the mid-section) were allowed to fix in formalin for greater than or equal to (≥) 48 hours prior to histological processing. Following routine processing, gonads were embedded in paraffin, sectioned at 5 micrometers (µm), and stained with hematoxylin and eosin (Luna, 1992). Reproductive stage was analyzed according to Blazer (2002) and given a score of 1–5 (figs. 2 and 3).
Images showing the reproductive stages in female Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish). A, Stage 1 is considered an immature fish and consists of dark basophilic, primary oocytes (an egg before maturation; indicated by arrows). B, Stage 2 consists of primary oocytes (dark basophilic) and early-stage cortical alveolar oocytes containing cortical alveoli (secretory organelles found along the periphery of the cytoplasm of mature oocytes, indicated by arrows) along the periphery. This stage is often observed during recrudescence, return of spawning after a period of abatement, in mature individuals. C, Stage 3 contains late-stage cortical alveolar oocytes (indicated by arrows) with the start of vitellogenin (egg-yolk protein) incorporation (early pre-spawn). D, Stage 4 contains mature, late pre-spawn vitellogenic (yolk forming) oocytes. E, Stage 5 is considered the post-spawn stage with few remaining vitellogenic oocytes (indicated by arrows), empty follicles (indicated by arrowheads) and degenerating/atretic oocytes (indicated by *), which will be reabsorbed for energy.
Histological images showing the reproductive stages in male Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish). A, Stage 1 is considered an immature fish and only consists of spermatogonia (immature germ cells [indicated by arrows]). B, Stage 2 consists of spermatogonia (indicated by arrows), dark eosinophilic, condensed spermatocytes (indicated by the letter a) and few spermatozoa within the lumen of the lobules. This stage is often observed during recrudescence, return of spawning after a period of abatement, in mature individuals. C, Stage 3 (early pre-spawn) contains fewer spermatogonia (indicated by arrow) and more spermatocytes (indicated by the letter a), spermatids (indicated by the letter b) and an increase in spermatozoa (indicated by the letter s). D, Stage 4 consists of lobules full of spermatozoa (indicated by the letter s) and is an individual close to spawning (late pre-spawn). E, Stage 5 is considered the post-spawn stage with few remaining spermatozoa (indicated by the letter s), distended lobules, and spermatozoa reabsorbed by Sertoli cells (indicated by arrows).
In brief, stage 1 individuals were sexually immature with early, primary oocytes (females; fig. 2A) or spermatogonia (males; fig. 3A). Stage 2 individuals were considered sexually mature, having testes containing spermatogonia, spermatocytes, and few spermatozoa in the lobule lumen (fig. 3B), and previtellogenic ovaries with the periphery of oocytes populated with cortical alveoli (fig. 2B). In both sexes, stage 2 was often observed during gonadal recrudescence. Stage 3 (early pre-spawn) testes contained few spermatogonia and an increasing number of spermatocytes, spermatids, and spermatozoa (fig. 3C) and ovaries were considered early vitellogenic with the first signs of vitellogenin globules incorporated into oocytes (fig. 2C). Stage 4 gonads were individuals that were late pre-spawn (figs. 2D and 3D) and close to spawning. In males this included testes containing predominately spermatozoa and in females it included ovaries with yolk-filled oocytes. Stage 5 in both sexes was considered post-spawn (post-ovulatory follicles and atretic oocytes in females; fig. 2E) and distended lobules mostly devoid of spermatozoa with reabsorbed spermatozoa in Sertoli cells in males (fig. 3E).
Results
Blue Catfish Collections
A total of 331 blue catfish were collected for this study, including 155 from Nanticoke River, 102 from Marshyhope Creek (Nanticoke River tributary), 53 from Patuxent River, and 21 from Broad Creek (Nanticoke River tributary; fig. 1). The number of each sex collected, mean total length (mm) and weight (g) are provided in table 1. All data is available in Walsh and others (2025).
Table 1.
Number of fish collected, sex, mean weight, and mean length of Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish) collected from tributaries of the Chesapeake Bay in Maryland and Delaware in 2020–22. Data are from Walsh and others (2025).[Date given in month/day/year format. n, number of fish collected; g, grams; mm, millimeters; M, male; F, female; ±, standard error of the mean]
Reproductive Endpoints
Histopathology
Adults were targeted for this study, but some of the fish collected were of adult size yet sexually immature (stage 1 individuals). The number of males and females in each reproductive stage at each site is provided in table 2.
Table 2.
Reproductive stages of male and female Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish) sampled from Marshyhope Creek, Nanticoke River, Patuxent River, and Broad Creek in 2020–22. Data are from Walsh and others (2025).To provide an overall examination of the seasonal changes and trends in reproductive stages of blue catfish, all samples were combined (fig. 4).
Bar graph showing the reproductive stage and number of female (F) and male (M) Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish) sampled from Marshyhope Creek, Nanticoke Creek, Broad Creek, and Patuxent River, 2020–22.
The number of female blue catfish with late pre-spawn (stage 4) vitellogenic ovaries started to increase in April (n=7) and was highest in May (n=12) followed by a decline in June (n=8) and July (n=3). There were no stage 4 females in August and one and three in September and October, respectively. During April, May, and June, there were no stage 5 (post-spawn) females; however, in July there were six. In males, the greatest number of late pre-spawn (stage 4) individuals was in April (n=19), followed by a subsequent decrease in May (n=15) and June (n=5). The number of late pre-spawn 4 males increased slightly again in July (n=8) and remained steady through October. The number of females with stage 2 ovaries was high in July–September, indicative of females who had spawned in the spring and were beginning gonadal recrudescence. In males, the number of stage 2 (n=3) and 3 (n=4) testes was highest in September but stayed relatively low or nonexistent throughout the rest of the year. There were three months when no stage 5 males were sampled, April, June, and October. Overall, these results demonstrate that blue catfish in these tributaries primarily spawn in late April–June, but a few remaining individuals spawn late in July.
Effective blue catfish management requires knowing the best time to harvest fish to remove the largest females to mitigate spawning. Females are typically capable of laying 4,000–8,000 eggs per kilogram (kg) of body weight (Chesapeake Bay Program, 2024), thus older and larger females could be a key target for removal. The largest stage 4 females were collected in April (n=7) and October (n=3). These results are similar to those of blue catfish sampled from the James and York Rivers in Virginia (Nepal and Fabrizio, 2021). The size of stage 4, female blue catfish (at Marshyhope Creek, Broad Creek, and Nanticoke River) are shown in figure 5 by the date they were collected. No stage 4 females were collected from the Patuxent River.
Graph showing the mean weight and total length of stage 4 (late pre-spawn), vitellogenic female Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish) collected from Marshyhope Creek, Broad Creek, and Nanticoke River in 2020–22.
Plasma Analyses
The results of the plasma analyses by month are provided in figures 6 and 7 and include the reproductive stage. Plasma was not collected from fish sampled from the Patuxent River. In both sexes, the total protein levels were inversely related to calcium (fig. 6).
Graphs showing the total protein and calcium concentrations in the plasma of female and male Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish) from Marshyhope Creek, Broad Creek, and Nanticoke River in 2020–22. A, females; B, males.
Calcium (Gillespie and de Peyster, 2004) and total protein (Ogŭz and Ünal, 2012) have been used in fish studies in tandem or as a surrogate for vitellogenin. Generally, these studies have documented a direct relationship between the concentrations of calcium and total protein, unlike the results here. The concentrations of calcium and total protein were similar between males and females throughout this study, and only in females did calcium levels respond similarly to E2, which was to be expected. It is possible that total protein was not a good alternative for vitellogenin in blue catfish.
In females, an increase in E2 was observed from March to April, as the spawning season commenced. E2 peaked in April and gradually decreased in May and June, then sharply dipped in July with the appearance of the first stage 5 (post-spawn) females (fig. 7).
Graphs showing the 17β-estradiol (E2) concentrations in the plasma of male and female Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish) from Marshyhope Creek, Broad Creek, and Nanticoke River in 2020–22. A, females; B, males.
This trend in females (a sharp increase in E2 at the start of the spawning season, a decline during spawning, and then a return to baseline levels) has also been observed in other fishes (MacKenzie and others, 1989; Johnson and others, 19919). E2 concentrations slightly increased in September and October, which coincided with the appearance of a few females with stage 4 ovaries, which is atypical during their recrudescence period. More sampling is needed to better understand why some females have vitellogenic ovaries in the fall. As there were also stage 4 males collected during this time, it will be important to determine whether blue catfish in this region are capable of batch spawning or if these observations are an example of some other type of spawning behavior.
In males, the highest E2 concentrations were detected in June with concentrations at or below 200 picograms per milliliter (pg/mL) the remainder of the year. In some species, an increase in E2 in males has been shown to be associated with the culmination of the spawning period (Chaves-Pozo and others, 2007). This increase in E2 in June also corresponds with the peak in total protein and dip in calcium. Since the trends in calcium and total protein were also observed in females, it is difficult to say how or whether they are specifically associated with E2 in males. However, collectively the results show that changes in the concentrations of these endpoints indicate that the spawning period began to wind down in June and fish transitioned into post-spawn reproductive patterns by July.
Conclusion
Management of an invasive species, especially Ictalurus furcatus (Valenciennes in Cuvier and Valenciennes, 1840; blue catfish), is a challenge, and understanding life-history traits is crucial for efficient mitigation strategies. The Chesapeake Bay Program’s Invasive Catfish Management Strategy states that one key to a successful management strategy of blue catfish is to understand their ecology and life history. To address this knowledge gap, the main goal of this study was to develop an assessment of the reproductive status of blue catfish for use across tributaries in the Chesapeake Bay region to enable practical comparison of reproductive and spawning synchronicity across geographic locations. The histology and E2 results from this study showed that blue catfish from tidal tributaries of the Chesapeake Bay in Maryland began to spawn in late April and peaked in May. By July, the appearance of stage 4 (late pre-spawn) females tapered off, E2 levels dropped, and multiple stage 5 (post-spawn) females were observed, indicating the end of the spawning season. Stage 4 (late pre-spawn) blue catfish of both sexes were sampled in fall, something that has also been observed in females of another catfish species. It remains unclear why this would occur, but potential reasons could include diapause, skipped spawning, year-round spawning, or overwintering for the next spawning season. To better understand this phenomenon, year-long sampling may help determine annual changes in sex steroid hormone levels and reproductive staging throughout the winter months.
The endpoints quantified in the plasma (E2, calcium, and total protein) were only collected from the Nanticoke River and its two tributaries, Marshyhope Creek and Broad Creek. Although multiple tributaries were included, the fish collected in Broad Creek and Marshyhope Creek may have originated from the Nanticoke River and vice versa; thus, were not geographically different populations and limited the scope of this study. In Virginia, a study of two Chesapeake Bay tributaries, the James and York Rivers (located in two different watersheds), had blue catfish populations with different reproductive traits attributed to differences in population densities and food and resource availability. Although this study did not compare river systems from different watersheds, it provided a better understanding of blue catfish reproduction in the Nanticoke River watershed. When compared to the findings of Nepal and Fabrizio (2021), blue catfish from the Nanticoke River began to spawn around the same time as fish from the James and York Rivers, but the largest females were found to spawn in late April. Nepal and Fabrizio (2021) found that blue catfish from the James River (which was more densely populated) not only matured at an older age and smaller size but also used more energy reserves for reproduction than fish in the York River. Other findings included differences in endpoints not examined in the current study, including gonadosomatic index, relative fecundity, egg organic content, and proportion of egg organic content. Males from Marshyhope Creek were larger than females, which was not always the case in the Nanticoke River, and reproductive differences may have been found had the same endpoints from Nepal and Fabrizio (2021) been analyzed. Thus, additional research in the tidal reaches of the Chesapeake Bay in Maryland could help to determine whether differences in reproductive characteristics exist, if they are associated with differences in population densities, and how that information could be applied to improve management practices.
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Glossary
- Cortical alveoli
Secretory organelles found along the periphery of the cytoplasm of mature oocytes.
- Fecundity
The number of eggs a female fish can lay during a spawning season.
- Lumen
Hollow center of the seminiferous tubules (coiled tubes in the testis that produce spermatozoa) in the testis, where spermatozoa are released.
- Oocyte
Female germ cell involved in reproduction.
- Sertoli cells
A type of somatic cell around which spermatids develop in the seminiferous tubules of the testis.
- Spermatogonia
Male, immature germ cell.
- Spermatocyte
Derived from spermatogonia and are the second stage in the formation of spermatozoa.
- Spermatid
Derived from spermatocytes and are the third stage in the formation of spermatozoa.
- Spermatozoa
Male sex cells that contain genetic material and are capable of fertilizing an egg.
- Spawning
Release or deposit of eggs and the fertilization with sperm.
- Recrudescence
A return of spawning after a period of abatement.
- Vitellogenesis
The process by which maturing oocytes accumulate yolk
Conversion Factors
Abbreviations
≥
greater than or equal to
≤
less than or equal to
BCA
bicinchoninic acid
E2
17β-estradiol; CAS ID: 50-28-2, Formula: C18H24O2, IUPAC ID: (17β)-estra-1,3,5(10)-triene-3,17-diol
EESC
Eastern Ecological Science Center
µL
microliter
µm
micrometer
mL
milliliter
n
number of samples collected
pg/mL
picograms per milliliter
USGS
U.S. Geological Survey
For more information concerning the research in this report, contact:
Center Director, USGS Eastern Ecological Science Center
U.S. Geological Survey
12100 Beech Forest Rd., Ste 4039
Laurel, MD 20708-4039
Or visit the Eastern Ecological Science Center website at
https://www.usgs.gov/centers/eesc
Publishing support provided by the Baltimore Publishing Service Center.
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Any use of trade, firm, or product names is for descriptive purposes only and does not imply endorsement by the U.S. Government.
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Suggested Citation
Walsh, H.L., Densmore, C.L., Regish, A.M., Norstog, J., Moore, J., Williams, B., Bressman, N., and Crum, Z., 2025, Reproductive parameters in invasive blue catfish (Ictalurus furcatus) from tributaries of the Chesapeake Bay in Maryland and Delaware, 2020–22: U.S. Geological Survey Open-File Report 2024–1074, 17 p., https://doi.org/10.3133/ofr20241074.
ISSN: 2331-1258 (online)
Study Area
| Publication type | Report |
|---|---|
| Publication Subtype | USGS Numbered Series |
| Title | Reproductive parameters in invasive blue catfish (Ictalurus furcatus) from tributaries of the Chesapeake Bay in Maryland and Delaware, 2020–22 |
| Series title | Open-File Report |
| Series number | 2024-1074 |
| DOI | 10.3133/ofr20241074 |
| Publication Date | January 31, 2025 |
| Year Published | 2025 |
| Language | English |
| Publisher | U.S. Geological Survey |
| Publisher location | Reston VA |
| Contributing office(s) | Eastern Ecological Science Center |
| Description | Report: vi, 17 p.; Data Release |
| Country | United States |
| State | Delaware, Maryland |
| Online Only (Y/N) | Y |
| Additional Online Files (Y/N) | N |