Laboratory‐bred Longfin Smelt produced offspring in the first year in captivity

Abstract

Objective

This study aims to develop the culture methods for Longfin Smelt Spirinchus thaleichthys in a laboratory setting, achieving the first production of a fully captive second generation (F2). The objective includes understanding the critical factors influencing the breeding and maturation of this species in captivity, contributing to conservation efforts and potential revival of this threatened species.

Methods

Longfin Smelt broodstock were collected from the San Francisco Estuary and reared in controlled conditions at the University of California Davis Fish Conservation and Culture Laboratory (FCCL). The study involved spawning of wild broodstock, rearing of first generation (F1) progeny to adulthood, and subsequent spawning to produce F2 offspring. Key variables such as salinity, temperature, and diet were carefully managed throughout the process. Fertilization and hatching rates were calculated, and the larvae were reared using the same methods to closely monitor and understand their growth and development.

Result

First generation Longfin Smelt successfully matured and spawned at age 1, earlier than commonly observed in the wild. From five crosses, four produced viable offspring. The study recorded a range of fertilization rates (10–40%) and high hatching rates (75–97%). Salinity was identified as a critical factor in both larval development and adult maturation. While initial larval survival was challenging, the larval rearing system used in this study proved effective.

Conclusion

The study represents a major breakthrough in the cultivation of Longfin Smelt, showing that it is possible to complete their life cycle under controlled conditions. It has provided valuable understanding of the maturation and breeding processes in these fish, with an emphasis on the influence of salinity. These outcomes are crucial for conservation strategies, offering the potential to help establish a refuge population and laying the groundwork for further research aimed at refining captive breeding methods for this species.

Impact statement

This research achieves a significant milestone in protecting Longfin Smelt, a threatened fish vital to both river and ocean environments. Successful breeding in labs takes us a step closer to their conservation, enhancing ecosystem health and aiding local fisheries. This work shows the power of dedicated conservation efforts in preserving at‐risk species and sustaining the balance of our natural habitats.

INTRODUCTION

The global decline in fish populations is a major concern, with many species becoming threatened or endangered due to various human activities (Arthington et al. 2016). The Longfin Smelt Spirinchus thaleichthys is one such species. This migratory forage fish is native to the west coast of North America, with the southernmost distinct population becoming increasingly at risk of extirpation (Hobbs et al. 2017; Sağlam et al. 2021). While the process of recovering imperiled fish species is complicated, conservation programs can use different strategies to revive populations, including habitat restoration, fisheries management, pollution and predation control, climate change adaptation, and captive breeding and rearing (Verhelst et al. 2021; Mitra et al. 2023).

Captive breeding and rearing programs not only play a vital role in increasing population size, preserving genetic diversity, and mitigating risks associated with threatened or endangered fish species (Eldridge and Killebrew 2008; Lessard et al. 2018), they also generate individuals suitable for both laboratory and field research, thereby contributing directly to conservation goals (Donaldson et al. 2019; Koch et al. 2022). However, despite studies demonstrating some Longfin Smelt aquaculture breakthroughs (Yanagitsuru et al. 2021, 2022; Mulvaney et al. 2022; Rahman et al. 2023), the successful production of the second generation (F2) in captivity from laboratory‐bred broodstock (F1) has never been achieved until recently. Here we describe the collection of wild adult Longfin Smelt broodstock, the maintenance and spawning of wild broodstock in a controlled environment, the rearing of F1 progeny to adulthood, and the production of the first fully captive F2 generation of Longfin Smelt.

METHODS

Between November 29, 2021, and March 18, 2022, a total of 375 Longfin Smelt broodstock were collected using trawl nets from low‐salinity habitats at two locations within the San Francisco Estuary (SFE): Alviso Marsh in South San Francisco Bay and Chipps Island near the confluence of the Sacramento and San Joaquin rivers in the upper estuary (hereon, “Delta”). The collection efforts were coordinated by a collaboration between the Fish Conservation and Culture Laboratory (FCCL) at the University of California, Davis (UC Davis), the UC Davis Otolith Geochemistry and Fish Ecology Laboratory (OGFL), and the U.S. Fish and Wildlife Service (USFWS). The broodstock were held in 20‐gal carboys and transported to the FCCL Byron, in California, USA, for rearing. These carboys were equipped with battery‐operated aerators and incorporated floating ice packs to maintain stable levels of dissolved oxygen and water temperature during transit. Upon arrival, the fish were quarantined and received a 3‐day prophylactic antibiotic treatment (Pennox 343, Animal Health International, Ceres, California, USA) before being further moved to rearing tanks in recirculating aquaculture systems, at a salinity of 10 ppt and temperature of 12°C. A total of 23 crosses were produced from the broodstock specifically collected on December 2, 2021, and March 2, 2022. Among them, only larvae from six crosses, all with USFWS broodstock from Chipps Island (Rahman et al. 2023; fork length: 105 ± 8.7 mm for females and 103 ± 6.0 mm for males), contributed to the data presented in this study.

From these six crosses, one was fertilized in filtered surface water from SFE that underwent treatment processes involving bead filters and ultraviolet (UV) disinfection with a salinity of 0.4 ppt while the remaining five were fertilized using 5 ppt saline water prepared with Instant Ocean sea salt (Spectrum Brands Inc., Blacksburg, Virginia, USA). The saline water was replaced with the same filtered surface water (0.4 ppt) after fertilization (about 5 min). The eggs were then incubated in 500‐mL plastic bowls for 6 days (1st step incubation: Rahman et al. 2023) and treated with Pond Rid‐Ich solution (55 mL Pond Rid‐Ich diluted with 378 mL water; Kordon LLC, Hayward, California, USA) for 1 min daily to reduce fungal growth (Tsai et al. 2021). On day 5, bentonite (Sigma‐Aldrich, St. Louis, Missouri, USA) was applied on the eggs to remove adhesion and coagulation (Tsai et al. 2022), and live and dead eggs were identified, separated, and estimated according to protocols developed for Delta Smelt Hypomesus transpacificus (Baskerville‐Bridges et al. 2005) at the FCCL on Day 6. The live eggs were moved into individual column incubators (2nd step incubation; Figure 1A). The fertilization rate (%) was estimated utilizing the protocols and formula described by Rahman et al. (2023). The water in the incubators was maintained at a salinity of 0.4 ppt and temperature of 12 ± 1°C until hatching (0 days posthatch [dph]; Table 1 and Figure 2). Daily maintenance was conducted on the incubators, and any dead embryos were recorded and removed.

Upon hatching, the larvae were transferred to randomly assigned tanks (133‐L black circular tanks with about 92 L working volume; Figure 1B) in one recirculating aquaculture system at 5 ppt and 12°C (Yanagitsuru et al. 2021) and reared for a duration of 40 days. The stocking density of larvae ranged from 10 to 31 individuals per liter. The larvae were nourished with rotifers (Brachionus plicatilis; L‐type, Reed Mariculture, Campbell, California, USA), along with newly hatched, unenriched Artemia nauplii (Artemia International, Fairview, Texas, USA). Other rearing conditions followed details described in Mulvaney et al. (2022). On 40 dph, the fish were counted and moved into 400‐L black circular tanks (about 320 L working volume; Figure 1C) in another recirculating aquaculture system at 5 ppt, 20 nephelometric turbidity units (NTU), and 12°C for another 40 days. The green water systems were established by adding commercial preserved algae (Nannochloropsis Instant Algae, Reed Mariculture; Hung and Piedrahita (2011)). Starting from 81 dph, the fish were supplemented with commercial dry feed (Biovita Starter mash crumble, Bio‐Oregon, Warrenton, Oregon, USA), and by 297 dph, the fish were fully weaned onto the dry feed (see Table 1). During the culture process, fish were consolidated into two 1100‐L black fiberglass tanks (about 860 L working volume; Figure 1D) due to space availability. At 386 dph, a total of 412 F1 fish were transported to the UC Davis Bodega Marine Laboratory (BML), Bodega Bay, California, USA, after being acclimated to the salinity at the BML by increasing 2 ppt every other day until 34 ppt. The fish were fed twice a day at the BML with a diet constituted with live adult Artemia and a dry feed mix to satiation. The dry feed mix included 35% freeze‐dried mysids, 30% freeze‐dried copepods, 30% Biovita Starter mash crumble, and 5% Spirulina powder.

RESULTS AND DISCUSSION

Shortly after their acclimation to 34 ppt (from 380 dph), the age‐1 captive male F1 Longfin Smelt started to show color changes (darkening) indicative of reproductive maturation, with additional fish showing similar signs of maturation after being transferred to seawater rearing tanks at the BML (Figure 1E). After 2 months in full‐strength seawater at the BML, several fish were expressing milt and eggs upon handling, indicating that they were ready to be spawned. A total of five crosses were subsequently made from these age‐1 mature F1 Longfin Smelt. Among them, four crosses generated viable offspring (spawner fork length: 90 ± 9.4 mm for females and 99 ± 10.2 mm for males). The first successful spawning of captive F1 Longfin Smelt was conducted on April 6, 2023, at 449 dph (Table 1; Figure 2), and another three crosses were made on May 17, 2023 (490 dph). The eggs were transported back to the FCCL (Cross 1) and to the UC Davis Academic Surge (Crosses 2–4) on the UC Davis campus for incubation. Cross 1 (egg number: 2700) had a fertilization rate of 30% and a hatching rate of 96%, and Crosses 2–4 (egg number: 4000, 3500, and 2200, respectively) yielded fertilization rates of 40%, 10%, and 30%, respectively, and hatching rates of 80%, 97%, and 75%, respectively. After hatching, the larvae in the UC Davis Academic Surge facility (Figure 1G) experienced mortality within a span of a few days. The larvae that hatched at the FCCL were subjected to rearing, utilizing a modified larval rearing system that had been previously established for Delta Smelt at the facility (Figure 1F; Tsai et al. 2022).

In the present study, we report the conditions used for culturing threatened Longfin Smelt F1 to adulthood and the production of F2 in the laboratory for the first time. The outcome demonstrates a huge step toward completing the life cycle in captivity, which will likely be crucial for supporting future conservation efforts, including the development of a refuge population program and the production of fish for supplementation and research purposes. The study also confirms that some Longfin Smelt may mature and spawn at age 1 (vs. age 2, as is commonly believed); however, it remains unknown whether this is a common occurrence in the wild population (Rosenfield and Baxter 2007; Lewis et al. 2020). The outcomes from this study indicate that salinity not only plays an important role in Longfin Smelt culture during their larval stage, as Yanagitsuru et al. (2022) reported, but also may act as a critical factor in their maturation, as no sign of maturation was observed during previous attempts of rearing F1 adults in freshwater at the FCCL. Future studies are warranted to determine the optimal salinities and stage‐specific transition timings that are needed to fully complete the Longfin Smelt life cycle in captivity and solidify the Longfin Smelt culture program as a valuable tool for conserving this imperiled species.

ACKNOWLEDGMENTS

The authors thank staff at the FCCL, the Academic Surge, and the BML for their extensive efforts toward managing rearing systems and caring for broodstock, F1, and F2 progeny. We also thank the field crews at the FCCL, the OGFL, and the USFWS for the collection and transport of wild broodstock. This study was funded by the California Department of Water Resources (#4600014181 to LSL, NAF, REC, and TCH) and the California Agricultural Experimental Station of the University of California, Davis (#CA‐D‐ASC‐2098 to NAF).

CONFLICT OF INTEREST STATEMENT

The authors declare that they have no conflicts of interest.

DATA AVAILABILITY STATEMENT

Data used for this article are available from the corresponding author upon reasonable request.

ETHICS STATEMENT

All animal care and handling were conducted in accordance with University of California–Davis International Animal Care and Use Committee requirements (Protocol #22969).

REFERENCES