Axenic and Gnotobiotic Aquatic Models: Methods, Husbandry, and Applications

1. Introduction

Microbial communities are integral components of animal biology regardless of the environment or species. In fish, microbes inhabit the skin, gills, and the digestive tract, as they do in the surrounding water. The effects can be beneficial, neutral or pathogenic depending on host developmental stage, environmental conditions and community composition. Descriptive microbiome studies have catalogued these communities in many species, but association alone does not establish whether a particular microorganism or microbial function is necessary or sufficient for a host phenotype. Axenic and gnotobiotic models address this limitation by placing microbial exposure under experimental control. In its strict experimental usage, an axenic or “germ-free” animal is maintained without detectable microorganisms, whereas a gnotobiotic animal has a microbiological status that is known, which may mean germ-free or colonized with one or more known microbial species. Aquatic gnotobiology therefore turns the microbiota from an uncontrolled environmental variable into a manipulable component of the experimental design [1,2].

Fish have several properties that make this approach highly practicable. In many teleosts, fertilization is external and embryos develop inside a chorion which is in direct contact with the water. The intact embryo can therefore be disinfected before hatching, after which larvae can be transferred into the sterile medium of choice. This avoids some of the invasive derivation procedures required in mammals and permits comparatively large cohorts to be generated from a single spawn. At the same time, the aquatic environment creates a distinctive complication: the animal and its water are microbiologically inseparable experimental compartments, thus any addition to the water, feed, inoculum, etc. can increase the contamination risk.

Larval transparency in zebrafish (Danio rerio) adds the ability to combine gnotobiology with live imaging, transgenic reporters and high-throughput genetic or chemical perturbations [3,4]. The field has consequently developed along two partly overlapping trajectories. The first uses zebrafish as a mechanistic vertebrate model to identify conserved microbial effects on epithelial maturation, immunity, metabolism and behavior. The second adapts gnotobiotic methods to aquaculture species in order to test probiotics, pathogens, live feeds and microbial management strategies under controlled conditions. These objectives require different husbandry solutions. A six-day-old zebrafish larva maintained in a tissue-culture flask is not equivalent to a first-feeding marine larva dependent on rotifers or Artemia, nor to a salmonid larva maintained for weeks before exogenous feeding. The derivation chemistry, water chemistry, feeding system and sterility-monitoring program must therefore be treated as species- and stage-specific parts of the experiment, not as interchangeable technical details [5,6].

This narrative review focuses on the practical production and use of axenic aquatic animals, with fishes as the primary emphasis. The review is based on the foundational zebrafish protocols, recent advances in longer-term germ-free culture, and representative models in medaka (Oryzias latipes), rainbow trout (Oncorhynchus mykiss), European sea bass (Dicentrarchus labrax) and Atlantic cod (Gadus morhua). Invertebrate systems are also included as they reveal principles that are directly relevant to aquatic gnotobiology. Attention was given to the points at which methodology can become a biological confounder: disinfectant exposure, antibiotic carry-over, diet sterilization, water-quality deterioration, low-biomass sterility testing and pseudo-replication within shared culture vessels. The aim of the present work is not to prescribe a universal protocol, but to define the experimental logic required to build and interpret a reliable axenic aquatic system.

2. Terminology and the Operational Meaning of “Axenic”

Terminology confusion can occur between “axenic” and “gnotobiotic” and the two terms should be used explicit because these labels are sometimes interchangeably although describe different experimental states. “Axenic” and “germ-free” is to be used strictly for animals in which microorganisms cannot be detected by the stated testing strategy. “Gnotobiotic” is, however, a broader term as it refers to animals whose microbial status is known, including axenic animals, monoassociated animals or animals colonized with defined multispecies consortia. “Conventionalized” usually describes an axenic animal deliberately exposed to a complex microbiota, often derived from conventional animals or their water. “Specific-pathogen-free” means that excluded pathogens have not been detected but it does not imply an absence of commensal microorganisms. Likewise, antibiotic-treated or microbiota-depleted animals should not be called “germ-free” unless the absence of detectable organisms has been demonstrated by appropriate assays [7,8] in sufficiently large number of replicates.

Even “axenic” is operational rather than metaphysical as no practical assay can prove the absolute absence of every bacterium, archaeon, fungus, protozoan or virus in every compartment of an animal and its culture system. Culture can miss viable but nonculturable organisms and broad-range PCR can miss targets below its detection limit. An experimental claim is therefore strongest when it is phrased as “no microorganisms detected under the stated culture and molecular testing conditions” and when those conditions are clearly described in a reproductible manner. This is especially important in aquatic systems, where a water sample is convenient but does not guarantee that the animal itself is sterile.

A useful conceptual distinction is between derivation, maintenance and validation. Derivation removes or excludes the microorganisms initially associated with the egg or embryo. Maintenance prevents reintroduction while preserving suitable animal physiology. Validation asks whether the desired microbiological state was actually achieved and retained, and thus failure in any one of these components invalidates an axenic interpretation. The most convincing gnotobiotic experiments add a fourth component represented by controlled recolonization, because restoration of a phenotype after defined microbial exposure provides causal information that a germ-free versus conventional comparison alone cannot supply [9,10].

3. Established Aquatic Gnotobiotic Models

Zebrafish is the most mature fish gnotobiotic system to date [9,11,12]. Foundational work established that germ-free zebrafish can be generated reproducibly and then colonized with complex microbiota or defined isolates, allowing direct tests of host responses to microbial presence and identity [9]. Detailed protocols standardized embryo collection, antibiotic preincubation, povidone-iodine (known as polyvinylpyrrolidone-iodine; aka PVP-I) and hypochlorite exposure, sterile washing, culture-flask or isolator husbandry, microbial association and sterility monitoring [3,7]. The model is particularly useful from embryogenesis through larval development, when optical transparency and genetic reporters permit tissue-level imaging. Recent work using gamma-irradiated formulated feeds extended germ-free zebrafish culture to approximately eight weeks, demonstrating that the traditional early-larval boundary is technical rather than absolute, while also showing that long-term germ-free growth and physiology require careful nutritional interpretation [13].

Medaka provides a complementary small-fish model with extensive genetics and inbred lines. A recent germ-free larval protocol adapted zebrafish-style embryo treatment to later-stage medaka embryos and demonstrated that microbial absence impairs aspects of intestinal epithelial maturation and defense-gene expression [14]. Medaka is therefore also valuable because its developmental timing, reproductive biology and genetic resources differ from zebrafish. The replication of microbiota-dependent phenotypes in another teleost helps distinguish broadly conserved fish biology simply by comparation.

Aquaculture fish species, unlike well researched laboratory models, impose more demanding husbandry constraints but provide greater ecological and translational relevance.

In rainbow trout, germ-free larvae can be maintained in the early beginning through yolk-dependent development and then fed sterilized feed. When gamma sterilization is not possible, microbiological controls should be performed. A gnotobiotic trout model showed that germ-free larvae were highly susceptible to Flavobacterium columnare and that recolonization with components of the endogenous microbiota restored resistance, including protection attributable to a commensal Flavobacterium isolate [6]. This is an important demonstration of colonization resistance in a farmed fish using a microbiologically controlled host rather than an indirect probiotic screening assay.

European sea bass gnotobiotic systems address another major aquaculture problem in terms of feeding: the dependence of marine larvae on live feed. Egg disinfection followed by sterile rearing and an axenic Artemia feed chain enabled controlled bacterial challenges and developmental studies [15,16]. The same platform has been used to visualize early Vibrio anguillarum interactions with larval tissues and to test candidate probiotic effects [17,18]. In parallel, Atlantic cod represents a marine model in which repeated glutaraldehyde treatment of eggs, antibiotic exposure before hatching and axenic rotifer/algal culture were integrated into a first-feeding system without continuing antibiotics after hatch [5].

These mentioned fish models situate within a broader aquatic gnotobiotic tradition, that includes crustaceans, rotifers, molluscs and other taxa [1]. In example, the water flee (Daphnia magna) is especially useful for ecological and nutritional questions because clonal hosts, defined diets and microbiota transfer can be manipulated simultaneously. Bacteria-free Daphnia display impaired growth, survival and reproduction, and microbiota effects depend on food availability [19,20]. Gnotobiotic Artemia is also widely used as a tractable host for probiotic and Vibrio-challenge experiments, and can serve as both an experimental organism and a controlled live feed in marine larviculture [21]. These models indicate that the central experimental advantage of aquatic gnotobiology is not sterility itself, but the ability to move reproducibly from “no detectable microbiota” to defined microbial complexity.

Table 1. Representative axenic/gnotobiotic aquatic models and their principal experimental strengths.

Model species	Axenic/gnotobiotic feature	Typical application	Reference
Zebrafish	Surface-disinfected embryos; flask or isolator culture; defined colonization; recent extension to ~55 dpf with irradiated feed	Development, immunity, metabolism, imaging, toxicology, microbial causality	[3,13]
Medaka	Embryo antibiotic pretreatment plus PVP-I/hypochlorite; sterile larval culture	Comparative teleost intestinal and immune development	[14]
Rainbow trout	Disinfected eyed eggs; long yolk-sac phase; sterile feed after first feeding	Colonization resistance, pathogen challenge, probiotic discovery	[6]
European sea bass	Disinfected eggs; sterile marine culture; axenic Artemia feed chain	Larval development, Vibrio challenge, probiotics	[15]
Atlantic cod	Glutaraldehyde egg disinfection; sterile seawater; axenic rotifer/algal feed chain	Marine first-feeding biology and host–microbe interaction	[5]
Daphnia magna	Bacteria-free clonal animals with controlled recolonization and diet	Fitness, nutrition, ecology, toxin tolerance	[19]
Artemia spp.	Axenic nauplii readily associated with defined bacteria	Probiotic mechanisms, Vibrio challenge, controlled live feed	[21]

Table 1. Representative axenic/gnotobiotic aquatic models and their principal experimental strengths.

Model species	Axenic/gnotobiotic feature	Typical application	Reference
Zebrafish	Surface-disinfected embryos; flask or isolator culture; defined colonization; recent extension to ~55 dpf with irradiated feed	Development, immunity, metabolism, imaging, toxicology, microbial causality	[3,13]
Medaka	Embryo antibiotic pretreatment plus PVP-I/hypochlorite; sterile larval culture	Comparative teleost intestinal and immune development	[14]
Rainbow trout	Disinfected eyed eggs; long yolk-sac phase; sterile feed after first feeding	Colonization resistance, pathogen challenge, probiotic discovery	[6]
European sea bass	Disinfected eggs; sterile marine culture; axenic Artemia feed chain	Larval development, Vibrio challenge, probiotics	[15]
Atlantic cod	Glutaraldehyde egg disinfection; sterile seawater; axenic rotifer/algal feed chain	Marine first-feeding biology and host–microbe interaction	[5]
Daphnia magna	Bacteria-free clonal animals with controlled recolonization and diet	Fitness, nutrition, ecology, toxin tolerance	[19]
Artemia spp.	Axenic nauplii readily associated with defined bacteria	Probiotic mechanisms, Vibrio challenge, controlled live feed	[21]

4. Generating Axenic Fish: Derivation as a Controlled Sequence

Successful axenic fish production usually begins before the disinfection step. The quality of embryos used, developmental stage, chorion integrity, broodstock health and the microbial load of the source water influence the margin between effective decontamination and chemical injury. Embryos of poor health tend to tolerate surface disinfectants less well, and those that are microbiologically heavily contaminated decrease the probability of mild protocols to succeed. As such, derivation should be validated on the specific fish line, facility water and developmental stage being used rather than treated as a fixed recipe. The objective is to reduce external microbial load as the chorion remains an effective physical barrier, before the animal begins feeding or extensively interacting with the surrounding medium [3,7].

The best characterized zebrafish protocols use a staged approach, embryos are collected cleanly, often incubated for several hours in embryo medium containing antibacterial and antifungal agents, washed, exposed briefly to approximately 0.1% PVP-I, washed again, and then exposed to dilute sodium hypochlorite before extensive sterile rinsing and transfer to sterile embryo medium.

A widely used culture-flask procedure employs the use of 0.1% PVP-I for 2 min followed by 0.003% sodium hypochlorite for approximately 20 min; exact timing and handling vary among established protocols [3,22]. These concentrations should not be generalized without validation. PVP-I and hypochlorite activity is affected by stock age, organic load and preparation, while embryo sensitivity changes with developmental stage. The critical experimental variable is therefore effective exposure and not merely nominal concentration.

A closely related protocol has been adapted for medaka. In one validated system, six-day-post-fertilization embryos were incubated for 4 h in embryo culture medium containing ampicillin, kanamycin and amphotericin B, rinsed and then treated with 0.1% PVP-I for 2 min, washed again, and then exposed to 0.003% sodium hypochlorite for 30 min before transfer to sterile culture flasks [14]. The longer embryonic period and timing of medaka hatching shows why a procedure should be adapted around developmental biology rather than copied solely on the basis of taxonomic similarity.

In the rainbow trout model, eyed eggs were rinsed, exposed for 24 h at 16° C to antibiotic/antifungal treatment, rinsed, treated with 0.005% sodium hypochlorite for 15 min, rinsed again and exposed to a 10 ppm iodophor for 10 min. After hatching, larvae were transferred to vented culture flasks containing sterile water without antibiotics [6]. This protocol exploited the long yolk-sac period as trout larvae can remain unfed while sterility is repeatedly assessed, postponing the difficult problem of sterile exogenous nutrition until approximately three weeks post-hatch.

Marine fish derivation often relies on glutaraldehyde, but here too the exact method is species-specific. In European sea bass, eggs have been disinfected with 200 mg/L glutaraldehyde for 3 min as part of a gnotobiotic rearing system, and bacteria-free status was assessed by both culture and bacterial 16S rRNA PCR [16]. In Atlantic cod, a protocol developed for reproducible bacteria-free larvae used two glutaraldehyde disinfection rounds separated by approximately one hour, followed by incubation of eggs in sterile seawater containing ampicillin and rifampicin until hatching; one treatment round was insufficiently reliable in that study [5]. Therefore, the most important lesson is that “stronger” sterilization is not automatically better. Over-treatment may reduce hatch, injure epithelial surfaces, alter developmental trajectories or even create later susceptibility that is incorrectly attributed to microbial absence.

Sterile water or any other medium must be prepared carefully. Freshwater embryo media can be made from defined salts using sterile stocks or 0.2 µm membrane filtration where appropriate. Seawater is more challenging because filtration, autoclaving and storage can change its chemistry [23]. Marine protocols often use filtered and autoclaved seawater prepared in advance [24]. Vessel selection should minimize unnecessary manipulations. Sterile tissue-culture flasks are practical for small fish larvae because they are closed, transparent and disposable; larger isolators or bottles are needed when feeding, aeration or longer culture periods increase exchange requirements. All components entering the system such as: water, air lines, feed, nets, tubing, anesthetic solutions, microbial inoculum and sampling tools must have a defined sterility or gnotobiotic status. Aseptic technique is therefore critical as a laboratory precondition. Transfers of specimens are best performed with sterile wide-bore pipettes or other low-trauma devices appropriate to fish size. The operator should plan the entire manipulation before opening the culture. The most common contamination events are mundane touching a nonsterile surface, reusing a reagent bottle, introducing unvalidated feed or sampling with tools that have not been sterilized prior use.

Before a new species, line or developmental stage is used for hypothesis testing, the derivation itself should be treated as a pilot experiment. The aim is to identify a window in which sterility success is high but hatching, morphology and early survival remain with an acceptable range. A practical pilot study can compare a small number of disinfectant exposure durations or concentrations while holding embryo stage, water and handling constant. Each condition should be evaluated for both microbiological outcome and animal outcome as selecting the treatment that produces the lowest microbial signal without considering developmental toxicity is a common mistake.

In zebrafish, established protocols provide a defensible starting point, but they also emphasize that reagent preparation, bleach activity and embryo handling influence success [3,7].

Once a local protocol has been selected, it should be kept for the main experiment rather than adjusted from batch to batch in response to early outcomes. Contamination tracing benefits from treating the culture workflow as a series of microbiological boundaries. The cleanest boundary is immediately after the final embryo wash, other risks include the sterile medium stock, transfer tools, vessel opening, feed, air supply and sampling. When contamination appears, testing retained aliquots of medium or feed and reviewing the timing of first positive surveillance can often distinguish derivation failure from later introduction.

Contamination detected immediately after derivation suggests inadequate egg decontamination or a compromised reagent, whereas a cluster of failures after first feeding points toward feed preparation or feeding technique. A contamination log should therefore record vessel identity, operator, feed batch, medium batch and every opening event. This level of traceability may appear excessive for a small larval study, but it is the fastest way to a stable gnotobiotic platform and is indispensable when cultures are maintained for weeks or even longer.

The transition from pilot derivation to an experimental run should also include predefined acceptance criteria. Useful criteria include a minimum hatching or early-survival rate, a maximum proportion of contaminated vessels, acceptable water-quality ranges and a rule for repeating an entire derivation when failure is systematic. These criteria prevent investigators from unconsciously selecting the most successful flasks after observing biological outcomes. Where embryo numbers permit, animals should be randomized among replicate vessels after the final sterile wash, and treatment assignment should be separated from the order in which vessels are handled. The same principle applies to colonization: inoculation order should not be confounded with treatment identity, because time outside the incubator or repeated opening of stocks can create systematic differences unrelated to the microorganism being tested.

Table 2. Examples of published fish derivation and early-rearing conditions. These are source-specific examples, not universal prescriptions.

Species	Representative derivation elements	Early maintenance / feeding	Key verification	Reference
Zebrafish	Antimicrobial preincubation; 0.1% PVP-I for 2 min; ~0.003% NaOCl (protocol-specific exposure)	~28–28.5°C; sterile embryo medium; small sterile flasks or isolator; sterile feed introduced according to protocol	Culture plus molecular testing; repeated surveillance	[3,7]
Medaka	At 6 dpf: 4 h ampicillin/kanamycin/amphotericin B; 0.1% PVP-I 2 min; 0.003% NaOCl 30 min	28°C sterile embryo culture medium; transfer to sterile flasks	Aerobic/anaerobic culture in published model	[14]
Rainbow trout	24 h antibiotic/antifungal exposure at 16°C; 0.005% bleach 15 min; 10 ppm iodophor 10 min	12 larvae/100 mL in vented flasks; yolk to ~20 dph; gamma-sterilized feed from 21 dph; partial water renewal	Multiple culture media plus 16S rRNA PCR on water and larvae	[6]
European sea bass	Published system includes 200 mg/L glutaraldehyde for 3 min with sterile seawater handling	Static/rotating sterile marine culture; axenic Artemia in feeding system	Marine culture/enrichment plus bacterial 16S PCR	[15,16]
Atlantic cod	Two glutaraldehyde disinfection rounds separated by ~1 h; eggs held in sterile seawater with ampicillin/rifampicin until hatch	Temperature increased gradually to ~12°C; axenic rotifer and algal feed system; no continued antibiotics after hatch	Prolonged culture-based surveillance of rearing and feed cultures	[5]

Table 2. Examples of published fish derivation and early-rearing conditions. These are source-specific examples, not universal prescriptions.

Species	Representative derivation elements	Early maintenance / feeding	Key verification	Reference
Zebrafish	Antimicrobial preincubation; 0.1% PVP-I for 2 min; ~0.003% NaOCl (protocol-specific exposure)	~28–28.5°C; sterile embryo medium; small sterile flasks or isolator; sterile feed introduced according to protocol	Culture plus molecular testing; repeated surveillance	[3,7]
Medaka	At 6 dpf: 4 h ampicillin/kanamycin/amphotericin B; 0.1% PVP-I 2 min; 0.003% NaOCl 30 min	28°C sterile embryo culture medium; transfer to sterile flasks	Aerobic/anaerobic culture in published model	[14]
Rainbow trout	24 h antibiotic/antifungal exposure at 16°C; 0.005% bleach 15 min; 10 ppm iodophor 10 min	12 larvae/100 mL in vented flasks; yolk to ~20 dph; gamma-sterilized feed from 21 dph; partial water renewal	Multiple culture media plus 16S rRNA PCR on water and larvae	[6]
European sea bass	Published system includes 200 mg/L glutaraldehyde for 3 min with sterile seawater handling	Static/rotating sterile marine culture; axenic Artemia in feeding system	Marine culture/enrichment plus bacterial 16S PCR	[15,16]
Atlantic cod	Two glutaraldehyde disinfection rounds separated by ~1 h; eggs held in sterile seawater with ampicillin/rifampicin until hatch	Temperature increased gradually to ~12°C; axenic rotifer and algal feed system; no continued antibiotics after hatch	Prolonged culture-based surveillance of rearing and feed cultures	[5]

5. Husbandry and Handling of Axenic Fish

Once derivation succeeds, husbandry becomes the main determinant of whether axenic status and animal health can be maintained. Conventional aquaculture systems depend heavily on microorganisms for decomposition and nitrification and an axenic system deliberately removes this biological buffering. Closed sterile cultures therefore accumulate ammonia, dissolved organic matter, uneaten feed and metabolic waste unless these are controlled by low stocking density, frequent medium exchange or special technological set ups. Oxygen limitation can also become acute because small and enclosed flasks have limited gas exchange, and because even microbial inoculation experiments themselves can consume oxygen. In monoassociation experiments with zebrafish, high bacterial inoculum concentrations caused mortality under some conditions in association with falling dissolved oxygen, because inoculum changes both microbial exposure and physical water quality [25].

Classical zebrafish larval culture is usually performed near standard developmental temperatures (approximately 28 – 28.5°C) with a defined light-dark cycle. Still, protocols differ in vessel size, fish density and medium-exchange schedule, but all depend on balancing sterility against sufficient water renewal [3,7]. The operator should record the actual number of animals per vessel and water volume rather than reporting only the gnotobiotic conditions, as specimen density affects oxygen availability, waste accumulation, microbial carrying capacity after colonization and the effective dose of waterborne compounds. It is therefore an experimental variable.

Feeding is the major transition in long-term axenic culture. Before exogenous feeding, embryos and yolk-sac larvae can be maintained with relatively few microbial inputs. Once feed is introduced, every particle is both a nutritional intervention and a potential source of contamination.

Autoclaving is a straightforward method, efficient and affordable, but alters vitamins, lipids, amino acids, and even texture and digestibility This is particularly problematic for small larvae with narrow nutritional margins. Gamma irradiation can sterilize dry formulated feed with less thermal damage, although irradiation may require validation for both microbiological efficacy and nutritional performance. In the long-term zebrafish study, gamma-irradiated feeds enabled germ-free culture to 55 days post-fertilization, with fish kept at approximately 28°C in small groups and subjected to repeated medium replacement and sterility testing [13]. The achievement is methodologically important, but smaller size and altered gene expression in long-term germ-free animals also emphasize that a “sterile diet” is not biologically neutral.

Rainbow trout has a different nutritional opportunity. Larvae can rely on yolk reserves for roughly the first 20 days after hatching, after which sterilized powdered feed can be introduced. In the published gnotobiotic trout system, gamma-sterilized food was provided every 48 h and half of the water volume was renewed shortly afterward to reduce accumulation of nitrogenous wastes and oxygen limitation [6]. This timing simplifies derivation but creates a clear methodological breakpoint because sterility risk and water-quality demand rise sharply at first feeding. Studies spanning this transition should treat pre-feeding and post-feeding periods separately in their quality-control plan.

Marine larvae add a further layer because many species require motile live prey. A gnotobiotic fish culture is only as controlled as its feed chain. European sea bass models have used axenic Artemia from approximately one week after hatching, while Atlantic cod protocols have combined bacteria-free rotifers with axenic algal cultures [5,15]. Live-feed cultures need independent microbiological testing because they can amplify a contaminant rapidly before introduction to fish. The feed should therefore be considered a separate gnotobiotic culture with its own batch identity, sterility records and failure criteria.

Handling procedures should be redesigned around asepsis rather than used unchanged from conventional husbandry. For imaging or sampling, animals can be transferred in sterile medium into sterile dishes or chambers and returned only if the procedure and all reagents have been validated. Anesthetic solutions should be prepared sterile or passed through a suitable sterile syringe filter if chemical properties permit and the container used for anesthesia should not be shared between microbiological treatments. Instruments used for dissection or euthanasia should be sterile and specific. When a terminal sample is needed for sterility validation, whole-animal homogenate is often more informative than water alone. For nonterminal work, water sampling can provide longitudinal surveillance but should be interpreted as an indirect measure.

Daily observations are especially important because morbidity can precede detectable contamination. For example, dead larvae, retained chorions and feed debris should be removed aseptically when feasible because decomposition rapidly degrades water quality and can amplify a low-level contaminant. Unexplained differences among vessels may identify handling problems, local contamination or water-quality failure before molecular assays do. Last but not least, routine prophylactic antibiotics after derivation should be avoided unless they are an explicit treatment variable. Continuing antibiotics can mask contamination, and constrain which microorganisms can be introduced, which is against the principal advantage of a gnotobiotic system [5].

6. Verification of Axenic Status

Sterility testing is the evidentiary foundation of an axenic experiment. A single negative plate at the end of a study is insufficient because contamination may be transient, localized, slow-growing or introduced late. A robust program combines longitudinal sampling with an endpoint assessment and uses at least two orthogonal detection principles. The appropriate schedule depends on the duration and manipulation intensity of the experiment, but testing should be more frequent after high-risk events such as derivation, first feeding, medium exchange through new equipment, microbial inoculation and invasive handling [7].

Culture-based testing remains indispensable because it demonstrates viable organisms and can be highly sensitive when relatively large sample volumes are incubated. Its weakness is selectivity: no single medium, temperature or oxygen condition grows all microorganisms. Fish gnotobiotic studies have therefore used multiple media and incubation conditions appropriate to the species and suspected contaminants. The long-term zebrafish feeding study evaluated irradiated diet sterility using bacterial and fungal media under aerobic and anaerobic conditions and monitored germ-free culture water repeatedly [13]. Rainbow trout cultures were assessed using several agar media and bacterial 16S rRNA PCR, with both water and animal samples included after feeding began [6]. Marine fish protocols similarly combine enrichment or plating with molecular tests [15].

Broad-range PCR or qPCR complements culture by detecting microbial nucleic acids independent of growth. Bacterial and mycoplasma 16S rRNA gene assays are common, and fungal ITS assays can be added when fungal contamination is plausible. The assay should include extraction blanks, no-template controls and a positive control of known low concentration. Detection limits should be reported or at least experimentally characterized. A negative PCR result is not proof of sterility if inhibitors are present or the sample volume is too small. Conversely, a positive result does not necessarily indicate viable contamination because DNA from dead organisms may persist after disinfection or feed sterilization.

Amplicon sequencing or shotgun sequencing can provide taxonomic resolution but should not be treated as automatically more reliable in low-biomass samples. Reagent and laboratory contaminants can dominate sequence libraries when true microbial biomass is near zero. Sequencing therefore requires negative extraction controls and a pre-specified rule for distinguishing background signal from biological contamination. Microscopy can add direct evidence for larger cells, biofilms or fluorescently labeled inocula, but routine light microscopy is generally less sensitive than enrichment culture or molecular detection for sparse contaminants.

Sampling location is as important as assay choice. Water is easy to sample repeatedly and is an excellent sentinel in a well-mixed system, but it does not fully represent microorganisms adherent to the fish, retained in the gut, attached to feed particles or present in vessel biofilm. Ideally, validation includes rearing water, representative animals and feed; terminal experiments can additionally test vessel rinse or biofilm material when contamination is suspected. A culture should be considered a failed axenic experimental unit if a contaminant is detected according to a pre-defined criterion. Selectively excluding contaminated individuals from a contaminated shared flask is usually not defensible because all fish in the vessel have experienced the same uncontrolled microbial exposure.

The sensitivity of the sterility program should itself be considered a measurable property. Laboratories establishing a new model can challenge their culture and molecular assays with low numbers of representative aquatic bacteria or fungi to determine whether the chosen sample volume and incubation scheme reliably detects contamination. This is particularly useful for long experiments in which a few cells introduced during feeding could expand slowly. Sampling 50 microliters of water and sampling several milliliters are not equivalent claims of surveillance, even when the same agar is used. Likewise, an enrichment broth can detect organisms that would be missed by directly plating a small volume. Reporting sample volume and enrichment strategy therefore gives the reader information needed to judge the strength of a negative result.

A second useful distinction is between surveillance and certification. Surveillance tests are frequent, minimally invasive measurements—usually culture of water or small aliquots of medium—designed to detect emerging problems without sacrificing animals. Certification is the stronger endpoint assessment used to support the final axenic claim and should include the host itself whenever possible. A well-designed study may use water culture throughout, then homogenize representative animals at the endpoint for culture and broad-range PCR or qPCR. If the feed was a major exposure route, retained feed from the same sterilized batch should also be tested. This layered approach is more informative than repeatedly applying a single assay to the same compartment and makes the microbiological evidence proportional to the strength of the biological conclusion.

Table 3. Complementary approaches for verification of axenic status.

Method	What it demonstrates	Strengths	Major limitation	Recommended role
Enrichment / plating	Viable organisms capable of growth under test conditions	Sensitive, inexpensive, directly demonstrates viability	Misses organisms that do not grow under selected conditions	Routine longitudinal testing using several media/conditions
Broad-range PCR / qPCR	Microbial nucleic acid above assay detection limit	Culture-independent; rapid; can be quantitative	Can detect DNA from dead cells; inhibition and low sample volume cause false negatives	Orthogonal confirmation, ideally bacterial 16S plus fungal ITS where relevant
Amplicon sequencing	Taxonomic identity of amplified microbial DNA	High taxonomic resolution; useful for tracing contaminants	Reagent contamination dominates low-biomass samples	Problem-solving or endpoint characterization with extraction blanks
Microscopy	Directly visible cells, biofilm, or labeled inoculum	Spatial information; useful with fluorescent microbes	Low sensitivity for sparse contamination	Supportive rather than sole sterility test
Animal + water + feed sampling	Distribution of contamination across system compartments	Reduces false reassurance from water-only surveillance	Often requires terminal animal sampling	Preferred endpoint validation; combine with longitudinal water tests

Table 3. Complementary approaches for verification of axenic status.

Method	What it demonstrates	Strengths	Major limitation	Recommended role
Enrichment / plating	Viable organisms capable of growth under test conditions	Sensitive, inexpensive, directly demonstrates viability	Misses organisms that do not grow under selected conditions	Routine longitudinal testing using several media/conditions
Broad-range PCR / qPCR	Microbial nucleic acid above assay detection limit	Culture-independent; rapid; can be quantitative	Can detect DNA from dead cells; inhibition and low sample volume cause false negatives	Orthogonal confirmation, ideally bacterial 16S plus fungal ITS where relevant
Amplicon sequencing	Taxonomic identity of amplified microbial DNA	High taxonomic resolution; useful for tracing contaminants	Reagent contamination dominates low-biomass samples	Problem-solving or endpoint characterization with extraction blanks
Microscopy	Directly visible cells, biofilm, or labeled inoculum	Spatial information; useful with fluorescent microbes	Low sensitivity for sparse contamination	Supportive rather than sole sterility test
Animal + water + feed sampling	Distribution of contamination across system compartments	Reduces false reassurance from water-only surveillance	Often requires terminal animal sampling	Preferred endpoint validation; combine with longitudinal water tests

7. Biological Consequences of Axenic Rearing in Fish

The scientific value of germ-free fish comes from the biological differences that appear when microbial signals are removed and then restored. Early gnotobiotic zebrafish work identified a broad set of microbiota-regulated intestinal genes and showed that a subset of host responses is conserved between zebrafish and mice [9]. These findings established fish gnotobiology as more than a technical curiosity: microbial colonization participates in core vertebrate programs of epithelial maturation, nutrient processing and innate immunity. Importantly, “the microbiota” is not a single stimulus. Different bacterial species and bacterial products can restore distinct components of the germ-free phenotype, and community composition can be reshaped by the host environment after transplantation [10,26].

Intestinal development is one of the clearest examples. Germ-free zebrafish larvae show delayed or incomplete features of epithelial differentiation, including altered brush-border enzyme activity, immature glycan patterns, reduced numbers of some secretory cell types, altered macromolecule uptake and increased intestinal transit. Conventionalization or association with individual bacterial strains can reverse different subsets of these phenotypes, demonstrating that multiple microbial cues converge on gut maturation rather than operating through a single generic inflammatory pathway [10]. In medaka, germ-free larvae likewise display abnormal absorptive vacuoles in intestinal epithelial cells and reduced expression of defense-associated genes, supporting a broader role for colonization in teleost intestinal maturation [14].

Epithelial proliferation is also microbiota-responsive. In zebrafish, resident microbes increase intestinal epithelial cell proliferation through signaling that requires the innate immune adaptor MyD88 and interacts with Wnt/β-catenin pathways. Monoassociation with Aeromonas veronii is sufficient to increase proliferation in germ-free larvae, illustrating how a defined isolate can be used to move from a community-level effect to a mechanistic pathway [27]. Microbial colonization also induces spatially and temporally structured NF-κB activity in the digestive tract rather than simply producing a uniform inflammatory state [28].

Innate immune development provides a second major example. Newly hatched fish rely strongly on innate defenses before adaptive immunity is mature. In germ-free zebrafish, microbial colonization primes neutrophils and induces inflammatory and antiviral mediators, increasing resistance to viral challenge in a process linked to TLR/MyD88 signaling [29]. Later work showed that microbial colonization can also dampen signaling by reducing MyD88 expression, helping establish an immune setpoint rather than continuously amplifying inflammation [30]. These observations are not contradictory: colonization can simultaneously induce defensive maturation and engage negative regulatory mechanisms that prevent excessive response. Gnotobiotic models are especially valuable for separating these phases.

Community members differ greatly in functional potency, experiments with defined zebrafish bacterial isolates demonstrated that relative abundance alone does not predict the magnitude of the host neutrophil response; a numerically minor member can exert a disproportionate immunomodulatory effect through secreted products [31]. This has an important implication for microbiome interpretation in aquaculture: an organism that changes only modestly in relative abundance may still be biologically decisive, while a dominant taxon may contribute little to the phenotype being measured. Gnotobiotic reconstruction is one of the few practical ways to test these hypotheses directly.

Microbiota also affects nutrient handling. In zebrafish, colonization stimulates intestinal fatty-acid uptake and lipid-droplet formation in the intestinal epithelium and liver, with effects that depend on diet and bacterial identity [32]. Bile-salt metabolism offers a more chemically resolved example: microbial transformation of bile salts modifies signaling through the nuclear receptor Fxr and influences intestinal epithelial and metabolic programs [33]. These studies illustrate why diet must be considered part of the host–microbe system. If sterilization changes feed composition, an apparent “microbiota effect” can be amplified or obscured by altered substrate availability.

The microbiota–gut–brain axis has also been examined in larval zebrafish. Germ-free status alters locomotor and anxiety-related behavior and changes responses to acute stress, while microbial exposure can modify these phenotypes [34]. Behavioral outcomes are particularly sensitive to husbandry confounders because water chemistry, density, nutrition and handling all influence activity and stress. Strong behavioral gnotobiotic studies therefore require matched vessel conditions and, ideally, recolonization controls rather than simple germ-free/conventional comparisons.

Long-term germ-free culture introduces additional questions. The 2026 zebrafish study using irradiated formulated feeds demonstrated germ-free maintenance to 55 days post fertilisation but found reduced size and substantial gene-expression differences relative to microbiota-replete comparators [13]. Such results should not be interpreted as a simple catalog of microbial functions. Extended germ-free development changes nutritional ecology, immune exposure and perhaps energy allocation simultaneously. Long-term studies must therefore separate the effect of microbial absence from consequences of the derivation process, feed sterilization, culture format and life-long exclusion of environmental microbes. The technical ability to keep fish germ-free longer increases rather than decreases the need for rigorous controls.

8. Experimental Applications: From Mechanism to Aquaculture

The most informative use of an axenic host is usually not the germ-free condition alone but a sequence of controlled associations. Monoassociation tests whether one microbial strain is sufficient to produce a phenotype. Defined multispecies communities test interaction and redundancy. Conventionalization asks whether a complex microbiota can restore a germ-free phenotype. Microbiota transplantation can reveal host filtering and community assembly. In zebrafish, reciprocal transplantation between fish and mice showed that host habitat strongly reshapes the relative abundance of transplanted microbial lineages, even when broad phylogenetic membership is retained [26]. This warns against assuming that the composition of an inoculum is the composition ultimately experienced by the host.

Association experiments require explicit control of dose, timing and duration. Zebrafish exposed to different concentrations of defined bacteria at different developmental stages show different survival, colonization and host-response profiles; very high inocula can create water-quality stress as well as stronger microbial exposure [25]. The appropriate unit is therefore not merely colony forming units per milliliter at inoculation. Investigators should report the culture volume, fish density, inoculum viability, time since inoculation, whether medium was exchanged, and microbial abundance at the time of phenotype measurement. Without these data, identical nominal doses can represent very different biological exposures.

Aquaculture applications are particularly compelling because early life stages frequently experience high disease pressure before vaccination is practical. Gnotobiotic rainbow trout demonstrated that the endogenous microbiota can provide direct resistance to F. columnare and allowed the protective effect to be traced to defined commensal bacteria [6]. European sea bass systems have been used to establish reproducible bacterial challenge conditions, visualize pathogen contact with the developing gut and evaluate probiotic candidates [15,17,18]. The key advantage is causal resolution: survival can be attributed to a defined host–microbe–pathogen combination rather than to an uncontrolled background microbiota.

Gnotobiotic platforms are also useful for nutrition, immunostimulation and environmental toxicology. They can determine whether a dietary ingredient acts directly on the host or indirectly through microbial metabolism, and whether a pollutant phenotype requires microbial biotransformation. Zebrafish reviews have highlighted the potential of germ-free models for mechanistic environmental-health studies [35]. For aquaculture feed development, however, translation requires caution. A response in a simplified germ-free larva may identify a mechanism, but commercial production occurs in a dense microbial ecosystem. Promising defined microbes should therefore progress from gnotobiotic proof-of-principle to conventional challenge models and ultimately to production-relevant trials rather than being treated as ready-to-deploy probiotics.

9. Lessons from Other Axenic Aquatic Animals

Aquatic invertebrates broaden the questions that can be addressed and highlight principles that are sometimes less obvious in fish. In Daphnia magna, removal of bacteria reduces growth, survival and fecundity, while recolonization restores performance [19]. The strength of this mutualism depends on food availability [20], demonstrating experimentally that microbiota effects are conditional on environmental resources. This is directly relevant to fish larviculture, where microbial phenotypes can change with feed type, ration and developmental transition.

Daphnia also provides strong evidence for microbiota-mediated environmental tolerance. Germ-free animals receiving microbiota from different donor genotypes acquire corresponding differences in tolerance to toxic cyanobacteria, showing that microbial community composition can transfer an ecologically relevant trait [36]. Such experiments offer a useful conceptual model for fish studies of harmful algal blooms, xenobiotics and variable water quality: a host genotype effect may actually be mediated partly through genotype-dependent microbiota assembly.

Gnotobiotic Artemia has become a practical screening system for microbial disease-control mechanisms. In a Vibrio challenge model, a PHB-accumulating Bacillus strain improved Artemia survival in a dose-dependent manner, with stronger protection associated with higher bacterial PHB content [21]. Artemia is therefore valuable both as a mechanistic host and as a controllable live-feed vehicle. The broader lesson is that gnotobiotic complexity can be built as a chain: sterile or defined algae feed sterile rotifers or Artemia, which in turn feed gnotobiotic fish. Each link must be validated independently because contamination introduced early in the chain will propagate downstream.

10. Major Limitations and Sources of Experimental Bias

The strongest criticism of any germ-free experiment is that “microbial absence” is rarely the only difference between groups. Derivation exposes embryos to disinfectants and often antibiotics; conventional controls may not experience the same chemical history. Feed sterilization can alter nutrient availability; sterile cultures may have different water chemistry from microbially active systems; and repeated aseptic handling can change density, stress and feeding opportunity. These effects are not reasons to avoid gnotobiotic models, but they require designs that partition microbial status from procedural history. A useful hierarchy of controls is: conventionally reared animals, germ-free animals, and germ-free-derived animals that are conventionalized or recolonized. Where feasible, a derivation-matched microbiota-replete control is stronger than an unrelated conventional cohort because it shares the chemical and handling history [13].

Antibiotic exposure deserves special attention. Antibiotics used before surface disinfection can improve derivation success, but they may enter the embryo, alter mitochondrial or developmental pathways, and select for resistant contaminants. Continued antibiotic supplementation after hatching can suppress rather than eliminate microorganisms and makes subsequent colonization dependent on antimicrobial resistance. The Atlantic cod protocol explicitly sought to avoid continued antibiotic exposure after hatch for this reason [5,37]. If antibiotics are required, their concentrations, exposure duration and washout procedure should be reported and matched across the relevant controls.

Diet is a second major confounder. Autoclaved, irradiated and chemically sterilized feeds are not necessarily nutritionally equivalent to the untreated diet. Long-term germ-free fish may therefore show growth differences caused by microbial absence, modified feed quality or an interaction between the two. A practical control is to feed the same sterilized diet to microbiota-replete animals and confirm acceptable growth and survival before attributing differences to germ-free status. The 2026 long-term zebrafish work used this logic when evaluating irradiated feeds, but the remaining differences between long-term germ-free and microbiota-replete fish still require cautious interpretation [13,38].

Pseudo-replication is a frequent statistical problem in aquatic gnotobiology. Fish sharing a flask also share water chemistry, microbial exposure and contamination events. For microbial treatments delivered into the water, the vessel—not each individual fish—is usually the independent experimental unit unless exposure is genuinely individual. Measuring ten larvae from one flask does not create ten independent microbial replicates. Robust studies replicate treatments across multiple vessels and, preferably, across independent derivations or spawns. Analyses can then treat individual animals as nested observations when appropriate. This design principle is crucial because a single contaminated or poorly oxygenated flask can otherwise dominate a treatment result.

Low-biomass sterility testing introduces another bias: false reassurance from negative tests and false alarms from contaminating DNA. Culture should include media and conditions capable of detecting plausible bacterial and fungal contaminants, while PCR/sequencing should include extraction blanks and negative controls. The testing program must be defined before outcomes are known. If a vessel becomes contaminated, the response—exclude the entire vessel, terminate the experiment or reclassify it—should follow a pre-specified rule. Post hoc retention of selected animals based on favorable phenotypes undermines causal interpretation.

Finally, gnotobiotic findings may be developmentally narrow. Most fish studies focus on embryos and larvae because they are easier to maintain and because early development is experimentally valuable. The microbiota, immune system, gut architecture and diet all change substantially during juvenile and adult life. Results obtained at 5–10 dpf in zebrafish should not be generalized automatically to mature fish or to aquaculture species with different developmental timing. Longer-term culture methods are beginning to close this gap, but they introduce new nutritional and husbandry challenges rather than eliminating them.

11. Minimum Standards for Reproducible Axenic Aquatic-Animal Experiments

The field would benefit from reporting standards that make an axenic experiment reproducible without requiring access to the originating laboratory. At minimum, authors should identify the animal strain or stock, embryo age or developmental stage at derivation, source and pretreatment of water, exact disinfectants and antimicrobial agents, exposure times and temperatures, number and volume of washes, vessel type, stocking density, temperature, photoperiod, feeding schedule and method of feed sterilization. “Generated according to a published protocol” is not sufficient when small changes in bleach activity, embryo age, fish density or feed processing can materially affect survival and sterility [3,39,40].

Microbiological reporting should specify what was tested and include a minimal checklist (Table 4), as well as how often and by which methods. Culture media, incubation temperatures, aerobic or anaerobic conditions and incubation duration should also be stated. Molecular assays should report target region, primer identity or assay reference, sample volume, extraction method and relevant controls. Sequencing-based sterility claims should include reagent blanks and a criterion for background subtraction. Feed and live-feed cultures should have their own validation records. If a study uses the term “germ-free,” the evidence supporting that status should appear in the main methods or a clearly accessible supplementary section rather than being implied.

Experimental design should identify the independent unit. When fish share a microbial environment, treatment replication should be performed across vessels. Independent derivations or spawns are highly desirable because they test whether results survive day-to-day differences in egg quality and decontamination efficiency. Randomization of embryos among vessels after derivation, blinded phenotyping where practical, and pre-defined exclusion criteria further reduce bias. Survival and contamination should be reported for each vessel, not only as treatment averages.

For colonization studies, the inoculum itself requires characterization. Investigators should report strain identity, culture conditions, growth phase, viability estimate, administered concentration, culture volume and exposure duration. Where conclusions depend on stable colonization, microbial burden should be measured after inoculation rather than inferred from the starting dose. Defined communities should report the input ratio and the recovered composition [12]. Timing matters: the same bacterium can produce different outcomes when introduced before or after major developmental transitions [25].

A minimum causal design should include, where biologically and logistically possible, a germ-free group, a microbiota-replete comparator and a recolonized or conventionalized group. Monoassociation or defined-community groups can then test sufficiency [41]. For long-term or nutrition-sensitive experiments, the microbiota-replete comparator should receive the same sterilized diet and similar vessel conditions. These controls do more than strengthen statistical confidence; they determine what question the experiment can legitimately answer. Without recolonization, a difference may reflect the entire derivation-and-maintenance procedure but with successful recolonization, the inference that microbes contribute causally becomes much stronger [4,14].

Table 4. Minimum reporting checklist for axenic fish experiments.

No.	Reporting checklist item
1	Define axenic/germ-free operationally and state exactly how it was tested.
2	Report embryo stage, water preparation, disinfectant identity/concentration, exposure time, temperature, and wash steps.
3	Treat stocking density, vessel volume, medium-exchange schedule, and feed ration as experimental variables.
4	Validate sterilized dry feed or live-feed cultures independently before introduction to fish.
5	Use longitudinal sterility surveillance and an endpoint test; include water and, when possible, animals and feed.
6	Combine culture-based and molecular detection rather than relying on one negative assay.
7	Predefine what constitutes contamination and how a contaminated vessel will be handled analytically.
8	Use the culture vessel as the microbial experimental unit when fish share waterborne exposure.
9	Replicate across vessels and preferably across independent derivations/spawns.
10	Include recolonized/conventionalized controls when making causal claims about microbial absence.
11	For colonization experiments, report viable inoculum dose, exposure time, culture volume, and recovered microbial burden.
12	Avoid continued antibiotics unless they are an explicit experimental treatment.

Table 4. Minimum reporting checklist for axenic fish experiments.

No.	Reporting checklist item
1	Define axenic/germ-free operationally and state exactly how it was tested.
2	Report embryo stage, water preparation, disinfectant identity/concentration, exposure time, temperature, and wash steps.
3	Treat stocking density, vessel volume, medium-exchange schedule, and feed ration as experimental variables.
4	Validate sterilized dry feed or live-feed cultures independently before introduction to fish.
5	Use longitudinal sterility surveillance and an endpoint test; include water and, when possible, animals and feed.
6	Combine culture-based and molecular detection rather than relying on one negative assay.
7	Predefine what constitutes contamination and how a contaminated vessel will be handled analytically.
8	Use the culture vessel as the microbial experimental unit when fish share waterborne exposure.
9	Replicate across vessels and preferably across independent derivations/spawns.
10	Include recolonized/conventionalized controls when making causal claims about microbial absence.
11	For colonization experiments, report viable inoculum dose, exposure time, culture volume, and recovered microbial burden.
12	Avoid continued antibiotics unless they are an explicit experimental treatment.

12. Future Directions

Metabolomics is particularly promising because many host effects are mediated through microbial transformation of nutrients and signaling molecules, as illustrated by bile-salt–Fxr interactions in zebrafish [33].

The current priority can be considered not simply extending the survival rate but developing nutritionally validated, scalable sterile diets and culture systems in which water quality can be controlled without introducing an undefined microbial biofilter.

A particularly promising direction is the production of biologically valuable materials from axenic fish. Fish skin is an obvious example as Atlantic cod skin is already used clinically as a regenerative scaffold to treat burns. Products such as Kerecis fish-skin grafts [42] which are currently applied by medical professionals to deep partial-thickness and full-thickness burns and other complex wounds [43,44], have been developed. These grafts are represented by decontaminated fish skin and thus their production could benefit from axenic specimens along with other extracellular-matrix preparations and other regenerative biomaterials.

Fish serum may represent an equally important opportunity. Fish serum has already been investigated as a substitute for fetal bovine serum in mammalian cell culture, including CHO cells, where appropriately treated fish serum can support cell proliferation [45,46]. Fish sera is also considered an indispensable reagent especially in fish primary cultures, as FBS is phylogenetically distant and may result in low culture yield [47,48]. Serum collected aseptically from validated axenic fish would have the additional advantage of originating from animals without an associated microbial community, including mycoplasmas. This is relevant because mycoplasma contamination has been documented in fish cell cultures and can heavily compromise experimental and biotechnological applications [49]. Developing standardized axenic fish for serum production could therefore open new biotechnology applications and reinforce the present ones.

13. Conclusions

Axenic and gnotobiotic aquatic animals provide a uniquely powerful bridge between microbiome description and causal biology. Fish are particularly advantageous because embryos can often be surface-disinfected externally and larvae can be manipulated in sterile water at useful experimental scale. Zebrafish has supplied the most detailed methodological and mechanistic foundation, but medaka, rainbow trout, European sea bass and Atlantic cod demonstrate that the approach can be adapted to distinct developmental strategies and aquaculture questions. Studies in Daphnia and Artemia further show that the principles extend across aquatic animal phyla.

The major technical challenge is not producing a nominally sterile embryo; it is maintaining a physiologically credible animal while continuously demonstrating that the microbial state remains controlled. Water quality, sterile nutrition, live-feed chains, handling, vessel-level replication and orthogonal sterility testing are therefore central scientific variables. Derivation chemicals and diet sterilization must be treated as possible confounders, and the strongest experiments use recolonization to distinguish effects of microbial absence from effects of the procedure itself.

As the field advances toward long-term culture, synthetic communities and translational aquaculture applications, methodological transparency will determine the reliability of its conclusions. “Axenic” should be reported as a validated operational state, supported by explicit detection methods and contamination rules. When these standards are met, aquatic gnotobiology offers one of the clearest experimental routes for identifying how individual microbes, microbial communities and microbial metabolites shape the development, health and disease resistance of fish.