Because poor water quality is the most common cause of environmentally induced diseases, some way to assess water quality is essential. Inexpensive test kits are easy to use and provide reasonably accurate information. Professional aquaculturists or advanced tropical fish hobbyists should be encouraged to purchase and use their own water-testing equipment. Pet fish owners often rely on pet stores to do the water testing; however, many pet stores have very limited capabilities in this area, use less accurate tests, and may be unable to accurately interpret results.
Veterinarians practicing fish medicine should have a comprehensive understanding of the dynamics and management of water quality, including general guidelines for acceptable water quality parameters (see Table: “Normal” Reference Ranges for Routine Water Quality Analysis). In addition to temperature, basic parameters of water quality can be grouped into four major categories: dissolved gases, nitrogenous compounds, pH and carbonate compounds, and salinity. The significance of water quality parameters varies with the type of system, species, and stocking density. Low DO and high ammonia are the two water quality parameters most likely to directly kill fish. Water quality interactions are dynamic and complex. Indirect relationships can lead to toxicity from other parameters, such as the effect of rising pH on (increased) ammonia toxicity, or there can be indirect relationships between water quality and certain infectious agents. For example, low or inappropriate temperature can be associated with fungal diseases of fish. A classic example is Fusarium solani infection of bonnethead sharks, which has been managed by raising the environmental temperature to >80°F (27°C).
“Normal” Reference Ranges for Routine Water Quality Analysis
See table: Common Environmental Diseases of Fish for an overview of common environmental diseases of fish.
Common Environmental Diseases of Fish
Aquatic organisms are sensitive to a wide variety of toxicants, particularly chlorine and chloramine, which are common additives to city water. Chlorine is also used to disinfect tanks and equipment. Chloramine is a form of chlorine that has been stabilized by amination. When treated for removal of the chlorine molecule, ammonia is released into the system. Both compounds are highly toxic to fish, with adverse effects seen at chlorine concentrations as low as 0.02 mg/L and mortality at 0.04 mg/L. A simple colorimetric test is available to measure chlorine and chloramine in aquatic systems. No chlorine or chloramine should be detected at any time live animals are present. Water samples for chlorine testing should be tested onsite; however, if that is not possible, they may be transported in glass bottles. The chemical can be transient and difficult to detect, so a negative test result may not completely exclude some contamination in the system being evaluated.
To test for chlorine and/or chloramine, kits are available that measure both free and total chlorine. Free chlorine measures hypochlorous acid (HOCl) and the hypochlorite ion (OCl–), which is the active property in bleach. Total chlorine measures free chlorine plus chlorine that is tied up as chloramine. Water treated with chloramines alone will test negative for free chlorine but have high amounts of total chlorine detectable; therefore, testing for both is important. When chloramines are treated with sodium thiosulfate to eliminate the chlorine, ammonia is released into the system. In such an instance, repeated water changes (each of which requires dechlorination, releasing additional ammonia) can result in high ammonia levels that also may be toxic. A properly conditioned biofilter should be able to metabolize the ammonia as it is released, but a new or damaged bacterial bed will not be able to manage the influx of ammonia from repeated deamination of chloramines. This problem can be overcome by using a dechlorinator specifically designed to deal with chloramines by also binding the ammonia byproduct. Effective use of dechlorination products requires testing water for both free and total chlorine before and after use. Following label instructions on products sold from pet stores usually effectively removes these chemicals; however, in rare instances more chlorine/chloramine than expected may be present in municipal water supplies. Treatment of water supplies will vary, and boluses of chlorine or chloramine may be sent through water lines as part of maintenance protocols in some circumstances. Further, inaccurate calculation of the volume to be treated can also lead to poor performance or failure of dechlorination products.
Chronic exposure to sublethal concentrations of chlorine is a surprisingly frequent problem, even for experienced aquarists. Veterinarians should test water for chlorine (free and total) every time a sample is submitted from a tank that uses a municipal water supply as source water. Clinical indications of sublethal chlorine exposure are nonspecific but may include ragged fins, excess mucus on skin and gills, cloudy corneas, behavioral signs such as lethargy or irritation, and sometimes a history of low chlorine level and chronic mortality.
Other toxicants include hydrogen sulfide and heavy metals. Hydrogen sulfide usually is a problem in poorly maintained tanks in which the sediments are not cleaned frequently enough, allowing anoxic areas to develop. Cleaning or other disturbance of these areas can release hydrogen sulfide into the water column, resulting in acute and catastrophic mortality. Another common source of hydrogen sulfide is well water; if this is the case, a distinctive “rotten egg” smell can sometimes be detected. Hydrogen sulfide is volatile and transient, so unless a water sample is collected at the time of the problem, a confirmed diagnosis may not be possible. Acute mortality has been reported at concentrations of 0.5 mg/L, but any detectable hydrogen sulfide should be considered a significant problem.
Heavy metals in water can result in acute, or more often, chronic mortality. If household plumbing includes copper piping, some copper may leach into the water. If released in sufficient volume, this may cause a fish kill. Problems are most likely when water has been allowed to stand in pipes. A copper test of suspect water should confirm the problem. Solutions include running the water before it is placed into the aquarium, or special filtration (eg, activated carbon) to remove metals.
Zinc toxicity has been associated with use of stainless steel vessels to house fish. It has also been reported rarely from public exhibits in which coins were allowed to collect on the substrate or to be ingested by fish.
Of the dissolved gases, oxygen is the most important. In outdoor production ponds, photosynthesis by algae is the primary source of oxygen. A diurnal cycle is established, which coincides with photosynthetic activity. During daylight hours, when photosynthesis occurs, oxygen levels rise and CO2 levels fall. At night, respiration is the driving force, resulting in a decrease in DO and an increase in CO2. Most finfish thrive when the DO concentration is >5 mg/L. When DO is <5 mg/L, fish become stressed; depending on species, size, and duration of exposure, a fish kill may result. Cardinal signs of a fish kill caused by hypoxia include sudden, significant mortality, usually noticed early in the morning (when oxygen levels are lowest); often, large fish are affected more than small fish. Fish that are hypoxic often school near the surface of the water and may be seen trying to gulp air, a behavior referred to as “piping.” Differential diagnoses for piping includes low DO, high nitrite, and gill disease.
Although low DO is most common early in the morning in outdoor ponds, it can occur at any time. Other common causes of low DO in ponds are cloudy weather, death of an algal bloom, excessive feeding, overstocking, and pond turnover. Unrecognized overstocking can occur in koi ponds if fish spawn successfully and offspring survive and are retained in the system. Pond turnover is a common cause of catastrophic mortality in pond fish. It occurs most frequently in deep ponds (>6 ft) and involves a phenomenon referred to as stratification. Water at the bottom of the pond cools, and a temperature gradient, called a thermocline, develops between warm surface water and cool bottom water. The thermocline acts as a physical barrier between the surface water (epilimnion) and bottom water (hypolimnion). Because photosynthesis, and hence oxygen production, occurs at the surface, the hypolimnion becomes hypoxic and develops a biologic oxygen demand. When the pond is mixed, or “turns over,” the oxygen is removed as the biologic oxygen demand of the hypolimnion is satisfied. This sudden removal of oxygen can result in oxygen depletion and a fish kill. The most common cause of pond turnover in the southern USA is a summer thunderstorm, in which energy released from cold rain coupled with wind and wave action is sufficient to mix the pond. Fish kills in Florida have occurred after hurricanes and have been attributed to pond turnover. Pond turnover can also be caused by seining, aeration, or other management practices that result in mixing of the epilimnion and hypolimnion. Fish kills caused by pond turnover can be avoided by performing a weekly oxygen profile during periods of greatest risk (usually during hot, summer weather). If stratification is detected, the pond should be aerated or mixed to break down stratified layers before a significant oxygen demand layer can develop. Turnover events resulting in localized areas of low DO are common causes of wild fish kills during the summer in lakes, ponds, and even rivers in the southern USA. Although rare, stratification-related phenomenon can occur in aquariums and other aquatic systems. Under some conditions, flow rate, current patterns (related to tank design), and oxygen demand can cause layering (ie, stratification) and consequent focal areas of anoxia.
When assessing DO and aeration in indoor systems or exhibits in which the primary source of DO is an aeration device, and water is clear, the percent saturation should be considered along with the total DO reading. The amount of oxygen that water can hold in saturation varies with water temperature, salinity, and altitude. Of these three factors, water temperature is the most important. As any of these variables increase, the amount of oxygen in solution at saturation decreases. Saturation tables are available to determine percent saturation for a given DO if temperature, salinity, and altitude are known. Many modestly priced oxygen meters now provide data on the concentration of DO (mg/L) as well as the percent saturation. If oxygen saturation is <100%, it may indicate inadequate aeration for the bioload or sanitation problems (development of anoxic, organic-rich areas within the system). In either case, an inability to maintain a system at, or very near, 100% oxygen saturation requires correction. Most fish do well if DO is >5 mg/L; however, the percent saturation should be considered an indicator of the system’s health.
Gas bubble disease is caused by supersaturation of water with dissolved gases. Although oxygen and/or CO2 can contribute to supersaturation, the predominant gas contributing to the problem is usually nitrogen. Gas bubble disease can result in acute catastrophic or chronic mortality. It may occur transiently and can be difficult to confirm. Supersaturation should be considered when unexplained mortality is encountered in an aquarium setting. One common source of supersaturation is the use of well water that contains high concentrations of nitrogen (gas) or CO2. This problem is easily remedied by aerating the water before it comes into contact with the fish. Common causes of gas bubble disease in public aquaria include the use of cavitating pumps, leaks in plumbing on the intake side (allow for gas to enter and be forced under pressure through the pump), and sometimes excessive turbulence in cold water exhibits. In these cases, the supersaturation is caused by atmospheric nitrogen gas. Gas bubble disease is manifest by exophthalmos and the presence of tiny gas emboli within fins, corneas, or other tissue. The presence of gas emboli within gill capillaries is diagnostic. Treatment of gas bubble disease is vigorous aeration to volatilize excess gas and correction of underlying mechanical problems. Confirming a case of supersaturation can be extremely difficult, especially if mortality was acute and gas emboli cannot be detected in tissue. Sometimes, tiny gas bubbles may be visible on the inside of the glass in an aquarium, suggesting a lot of gas is in the water column. A saturometer will measure all dissolved gases and is the best tool for direct detection of the condition. If DO of the system is known, this equipment can be used to calculate the concentration of nitrogen gas present. Permanent correction of the problem includes identification and correction of the source of the excess gas.
CO2 can be toxic to fish at concentrations >12 mg/L. The concentration of CO2 in solution in ground water is typically <10 mg/L. Water from affected systems often is acidic (pH <7). A quick field test for excessive CO2 involves vigorous aeration of a bucket of suspect water for 1 hr. A significant increase in pH (ie, >1 unit) over the hour is indicative of excess CO2. Fish exposed to high concentrations of CO2 may be quite lethargic and even disoriented. Hybrid striped bass exposed to toxic levels of CO2 (~40 mg/L) at the surface have been observed with their backs out of the water. These fish reacted dramatically to salt added to the affected tank by trying to jump out of the water. When CO2 is high in the water column, fish are not able to release it from the bloodstream, resulting in hypercarbia and acidemia. The condition is exacerbated by low concentrations of DO. Nephrocalcinosis and visceral granuloma were reported in salmonids exposed to a high level of CO2 in the water, leading to metabolic acidosis and urinary and tissue precipitation of calcium, around which extensive granulomas developed. Treatment for CO2 toxicity is increased and vigorous aeration. Stocking density should be assessed and may need to be decreased.
Nitrogenous wastes enter the aquatic system directly from excretion by fish, decomposition of organisms in the water, or degradation of fish food. Fish foods are generally very high in protein, often >38%, and can add significant quantities of nitrogen to a system. Nitrogen is eliminated from fish by passive diffusion of ammonia (NH3) from gill capillaries. Once NH3 is released into the water, it enters the nitrogen cycle, a natural process in which bacterial populations change ammonia to nitrite (NO2) and then to nitrate (NO3). Nitrate is most commonly removed from aquariums by water changes. In large, commercial systems, discharge of salt water to municipal water supplies is not allowed, and nitrate accumulates. It can be removed by anaerobic filtration, which converts NO3 to nitrogen gas (N2), which is volatile and quickly leaves the system. These anaerobic filters are expensive and can be challenging from the design perspective, so they are not common except in very large exhibits. Plants or algae, if present in a system, will use nitrogen products directly. Toxicity of each of these parameters is discussed below.
NH3 is highly toxic and frequently limits fish production in intensive systems. It is also dynamic, and when it enters the aquatic system, an equilibrium is established between NH3 and ammonium (NH4+). Of the two, NH3 is far more toxic to fish, and its formation is favored by high pH (>7) and water temperature. When pH exceeds ~8.5, any NH3 present can be dangerous. In general, a normally functioning aquatic system should contain no measurable NH3 because as soon as it enters the system, it should be removed by aerobic bacteria in the environment. Ammonia test kits do not typically measure NH3 directly but instead measure the combination of NH3 and NH4+, referred to as total ammonia nitrogen (TAN). A TAN <1 mg/L is usually not cause for concern unless the pH is >8.5. However, if the amount of NH3 is increased, an explanation should be sought. The amount of toxic NH3 present can be calculated using the TAN, pH, and water temperature. When NH3 levels exceed 0.05 mg/L, damage to gills becomes apparent; levels of 2 mg/L are lethal for many fish. Fish exposed to ammonia may be lethargic and have poor appetites. Acute toxicity may be suggested by neurologic signs such as spinning, disorientation, and convulsions.
Overfeeding or malfunction (death) of a biologic filter are common causes of increased NH3. If possible, a water change (≥50%) should be done as soon as high NH3 levels are detected. When changing water to alleviate NH3 toxicity, it is imperative to consider whether source water contains chloramines, because this can contribute to increased NH3 concentrations. If TAN is extremely high (ie, >5 mg/L) and pH is acidic (ie, <6), fish should be moved to a clean system (tempered for pH and temperature) to avoid a sudden shift from NH4+ to NH3 as the pH rises during the water change. Feeding should be discontinued or significantly reduced until the problem has been corrected.
Two conditions encountered in pet fish medicine are characterized by high NH3 concentrations, appropriately called new tank syndrome and old tank syndrome. New tank syndrome occurs when NH3 levels rise during the first 2–3 wk after a new system is set up, because the biofilter has not had time to develop. In this situation, the NH3 concentration will be increased, but all other parameters should be within normal limits. Beginning aquarists are likely to overstock and overfeed new systems, resulting in significant NH3 spikes and, subsequently, sick or dying fish. Daily monitoring of TAN coupled with frequent water changes to manage NH3 will be necessary until the biofilter cycles. Maturation of the biofilter will be indicated by decreasing concentrations of TAN and increasing concentrations of NO2 (which will decrease as the filter conditioning process is completed). Damage to a biofilter can be caused by use of antibiotics or other chemicals and result in a similar situation. It usually takes ~6 wk for a new biofilter to cycle. When this time frame is extended, there may be complications attributed to poor design, use of chemicals, or lack of adequate oxygen and carbonate (alkalinity) in the filter bed. To prevent new tank syndrome, aquarists use several “tricks” to get biofilters started. These include purchasing commercially available nitrifying bacteria from a reputable source, “feeding” the bacteria with fish food or ammonium chloride before adding fish, or adding fish slowly to the new system.
Old tank syndrome is less frequently recognized. It is characterized by extremely high NH3 levels (TAN may be >20 mg/L), extremely low pH (usually <6, may be <5 in severe cases), and a complete absence of alkalinity. The condition is caused by complete exhaustion of buffering capacity within a system, usually precipitated by improper management over a period of months. Over time, the biofilter bacteria acidify the water through the nitrification process. As the buffering capacity (alkalinity) is exhausted, organic acids that have accumulated drop the pH, and the acidic environment kills the biofilter, leading to an accumulation of NH3. When correcting such a situation, it is important to eliminate as much “bad” water as possible and avoid a shift in residual NH3-H to the toxic un-ionized state (NH3) as pH rises. A simple water change under such circumstances can result in catastrophic mortality as pH rises above 7 and ammonium shifts to un-ionized (toxic) ammonia. Over-the-counter products that chemically remove or bind NH3 can help prevent mortality, but the system must be thoroughly cleaned and restarted. It will take several weeks for the system to recover.
The second breakdown product in the nitrogen cycle is nitrite (NO2), which is also toxic to fish. Most test kits actually measure nitrite-nitrogen rather than nitrite. A conversion factor of 3.3 can be used to calculate the actual nitrite concentration (3.3 × mg/L NO2−N = mg/L NO2). NO2 can enter the bloodstream passively across the gill epithelium. It complexes with hemoglobin to form methemoglobin, resulting in methemoglobinemia, or brown blood disease. As in other species, RBCs containing methemoglobin are unable to transport oxygen, resulting in a physiologic hypoxia regardless of oxygen content in the water. There are species-specific differences in fishes’ susceptibility to NO2 toxicity (eg, centrarchids [bass, bluegill, etc] are refractory). Marine fish were thought to be protected from NO2 toxicity by salts in their environment; however, red drum have developed brown blood disease in the presence of NO2. A tentative diagnosis of brown blood disease can be made by observing the characteristic chocolate brown color of the gills, although this change is not detectable until methemoglobin levels are substantial. In severe cases, the color of blood samples will also be abnormally dark. Methemoglobin concentrations in the blood can be determined, although this is not necessary for clinical management. A water quality test can confirm the presence of NO2. Fish affected with methemoglobinemia typically show signs of hypoxia, often manifest by piping.
The most rapid treatment for NO2 toxicity is a water change, but this is not feasible in large ponds. Increasing chloride (Cl–) concentration in the water creates a competitive inhibition at the gill epithelium between Cl and NO2. Many ornamental ponds and aquaria are maintained with residual chloride levels because of the addition of salt (1–3 ppt) as a relatively permanent treatment. In these cases, nitrite is less likely to be a problem, because chloride levels are increased by the residual salt concentration. In freshwater production ponds for channel catfish, a ratio of 6 parts Cl to 1 part NO2 has effectively prevented or treated methemoglobinemia caused by nitrite exposure. The absorption of Cl– across the gill membrane reduces the amount of NO2– entering the bloodstream; consequently, the percentage of hemoglobin converted to methemoglobin is decreased, resulting in immediate relief to the fish and a cessation of mortality usually within 24 hr. To determine the amount of salt required to increase chloride levels in large ponds, the concentrations of NO2 and Cl present must be measured by commercial test kits. The concentration of Cl needed (mg/L) = (6 × NO2) − Cl present. Once the necessary concentration of Cl– to be added is known, the volume of water can be calculated in acre-feet or gallons (1 acre foot = 1 surface acre, 1 foot deep or 325,850 gal.), and salt can be added to achieve the desired Cl level (4.5 lb of salt will add Cl at 1 mg/L to 1 acre-foot of water, or 1 lb of salt will add 1 mg/L Cl to 72,411 gal.). In aquariums and garden ponds, a water change and filter maintenance are recommended to correct nitrite problems; however, salt may still be used to halt mortality during a sudden increase in NO2 exposure for many freshwater fish.
Although considered less toxic than ammonia or nitrite, chronic exposure to nitrate (NO3−N) has been associated with development of goiter in some species of elasmobranchs. As mentioned earlier for the nitrite test kit, most test kits for nitrate actually measure NO3−N. To convert to the actual NO3 concentration, this number must be multiplied by a correction factor of 4.4. Practitioners should read the literature carefully to distinguish reports of NO3-N versus NO3 concentrations. Chronic exposure to NO3−N concentrations of 70 mg/L resulted in histologic evidence of goiter in white spotted bamboo sharks within 29 days of exposure. Chronic exposure to NO3-N of 35 ± 5.12 mg/L resulted in development of overt goiter and 100% mortality of brown spotted bamboo sharks and 18% mortality of white spotted bamboo sharks in a recirculating system after ozone was added. Goiter is a complex disease, and there seem to be species-specific differences in susceptibility that are not well understood. Contributing factors include inadequate dietary iodine or environmental iodide, ozonation, and nitrate exposure. Goiter is characterized by a ventral midline swelling in the cervical region of elasmobranchs. Diagnosis can be confirmed by measuring T4. In healthy (no clinical or histologic evidence of goiter), captive, white spotted bamboo sharks housed in a natural seawater system, T4 was 14.77 ng/mL (range of 9.57–30.50 ng/mL in five animals). Lower levels have been reported in sharks with visual evidence of goiter. If an animal is necropsied, thyroid tissue can be evaluated histologically to confirm the diagnosis. Nitrate is well recognized as a goitrogenic compound and may be present in fairly high concentrations (>70 mg/L NO3-N) in recirculating aquarium and aquaculture systems. Nitrate blocks the uptake of iodine by the thyroid gland, resulting in an inability to produce thyroid hormone and constant stimulation of the glandular tissue. Fish and elasmobranchs absorb micronutrients, including iodine, from the water column. In an ozonated system, the problem is exacerbated because iodine is converted to iodate (IO3), which is not biologically available. Dietary supplementation of iodine at 10–30 mg/kg/wk is recommended for elasmobranchs to prevent development of goiter. Environmental iodine should be maintained with concentrations of 0.15 µM (0.01–0.02 mg/L) iodide (I–). Potassium iodide (Lugol's solution) has been used to increase environmental iodide concentrations in public aquaria. Goiter occurs in teleosts and other aquatic species but is not as easily recognized. The body shape of many elasmobranchs allows for easy recognition of the problem during a visual examination.
The carbonate cycle is an important concept in water quality management, and its complexity is reflected in the dynamic interactions between CO2, pH, total alkalinity, and total hardness. In aquatic systems containing algae or plants, CO2 fluctuates on a diurnal basis, similar but opposite to fluctuations in DO. As CO2 concentration changes, the pH of the water also changes. As CO2 concentration decreases during daylight hours, pH rises, peaking in late afternoon. Conversely, as CO2 concentration increases during the night, pH falls, reaching its lowest level just before daylight. A diurnal pH change from 6.5 to 9 is not unusual in a freshwater fish pond with a healthy algal bloom. Most freshwater fish can tolerate reasonable fluctuations in pH, and the lethal limits for many species are approximately 4 and 10. Marine fish are much less tolerant of pH fluctuations; the marine environment is much more stable, with a pH of 8.2–8.3. For marine tanks, a pH in the range of 7.8–8.5 is usually considered normal.
Although fish kills caused by improper pH are rare, hydrated lime (Ca[OH]2) is sometimes added to freshwater ponds by mistake. Ca(OH)2 will rapidly increase the pH to >10, killing all fish present. Correct liming of ponds is mentioned briefly below.
CO2 released into an aquatic system enters the carbonate cycle: H2O + CO2 ↔ H2CO3 ↔ H+ + HCO3– ↔ 2H+ + CO32–. The process is driven by the presence of carbonate (CO32–) in the system, which is measured by testing the total alkalinity (TA). For most fish, water should be of moderate alkalinity, 100–250 mg/L. When TA is <50 mg/L, water is considered low in alkalinity, and buffering ability will not be adequate to prevent major pH fluctuations. Toxicity of copper sulfate, an algicide and effective parasiticide, is closely associated with TA, and the compound cannot be used safely if TA is <50 mg/L. To raise alkalinity, dolomite (CaCO3 and MgCO3) or agricultural limestone (CaCO3) may be added to the system. Dolomite is most convenient for small systems and can be purchased in 50-lb bags and used to effect. Baking soda (NaHCO3) can also be used to increase alkalinity in small systems. To raise alkalinity in outdoor ponds, agricultural limestone is commonly used; the method is similar to “liming” a pasture. Soil samples can be tested to determine how much lime needs to be added, but in general 1–2 tons per surface acre works well. The limestone should be unloaded on the bank of the pond and then usually has to be moved into the water using a shovel and boat. It does not need to be distributed throughout the entire surface area of the pond, but it takes several weeks to get into solution. Consequently, the alkalinity will change slowly, so it should be monitored for several days, or weeks if necessary, after the addition of these compounds. Lack of alkalinity can impair biologic filtration, resulting in accumulation of ammonia in a system. Alkalinity should be ≥100 mg/L in freshwater systems and ≥250 mg/L in saltwater systems.
Total hardness (TH) should not be confused with TA. Both TH and TA are reported as mg/L of CaCO3. The difference is that the test for TA measures the HCO3–, OH–, and CO32– fraction, and the test for TH measures the calcium (Ca2+) fraction. The test for TH also measures other divalent cations in the system, including magnesium, manganese, iron, and zinc. TH is important in determining the amount of calcium available to young fish. Calcium chloride, dolomite, or agricultural limestone can be added to water to increase calcium concentration. For channel catfish, TH >20 mg/L is required for normal skeletal growth and development. Fish absorb minerals from the water column; therefore, use of water with very low TH, which can be caused by use of deionized water, can result in poor growth and mortality.
The salinity of seawater is determined by a complex array of salts. Seawater is ~3% salt, which is 30 ppt (30 g/L). For marine fish, many of the micronutrients present in seawater are essential, so it is necessary to buy or make “sea salts.” In freshwater, salinity may be increased using table or water softener salt (NaCl). Salt is often used in freshwater systems to reduce osmoregulatory stress or to eliminate certain ectoparasites. Salinity can be measured with a clinical refractometer or with a hydrometer purchased from a pet store. It is important not to confuse the chloride (Cl–) test, which is used in assessment of NO2:Cl in freshwater systems with high nitrite, with salinity. The chloride test measures ppm chloride; if any salinity at all is present in the water (ie, salt has been added), the test will not work properly because the amount of chloride present is so high. The easiest way to calculate the amount of salt needed to increase salinity is to calculate the total volume in liters (3.8 L = 1 gal.), remembering that 1 g/L = 1 ppt. Most non-pond freshwater systems can be maintained with a residual salinity of 1–3 ppt, whereas most saltwater systems will have a salinity of 30–33 ppt. Some freshwater species (eg, wild-caught Amazonian fish) may not tolerate permanent exposure to the low levels of salt mentioned above.
Environmental temperature is extremely important to the health and well-being of fish and other aquatic species. As poikilotherms, fish have a very limited ability to control body temperature, and physiologic systems are designed to work optimally at species-specific temperature ranges. Sudden changes in temperature, of even just a few degrees, can result in compromise of immunity and increased pathogenicity of some infectious agents. Some fish (eg, channel catfish or koi) are very tolerant of a wide range of environmental temperatures; however, this does not imply that drastic temperature fluctuations are acceptable even for these species. Others, such as discus, thrive in only a very narrow temperature window. When evaluating housing and husbandry, practitioners should know the temperature at which animals are being housed and confirm that it is appropriate for the species. Suboptimal environmental temperature is an important component in some fungal infections. Many infectious agents, especially viruses, have specific temperature windows at which they cause clinical disease and mortality. Fish infected but maintained at temperatures above or below these optimal ranges are more likely to survive infection but may become carriers. When handling or transporting fish, moderating temperature change is essential. A general rule is 1°F, or even 1°C, per hour as a maximum change. Some fish may tolerate more or less of a change over time.