How To Treat A Discus Fish With Formalin


How To Treat A Discus Fish With Formalin
At moderate water temperatures (less than 70(F or 21(C), fish can be left in a 250 mg/l formalin bath for about one hour ; however, if fish are weak or noticeably sick, the treatment should be discontinued after 30 minutes.

How do you use formalin for fish?

Application of Formalin as a Parasiticide for Fish – Formalin is added to the water as a bath treatment. The dosage rate used is determined by the period of time the fish are to be in contact with the chemical, and by the condition of the fish. At lower dosages, formalin can be applied as a prolonged bath, which means it is put directly into the system water with the fish and left indefinitely.

By keeping the fish in their original system for the treatment, the fish are not subjected to additional handling stress; pathogens are not spread to other areas; and parasites in the system water are exposed to the treatment, helping reduce reinfection. The concentration of formalin appropriate for a prolonged bath is 12.5 to 25 mg/l.

The higher concentration of 25 mg/l is easily applied to smaller systems by adding 1 milliliter (ml) of formalin per 10 gallons of system water (or 2 drops of formalin per gallon of water). The lower concentration of 12.5 mg/l would be appropriate for more sensitive fish species or extremely sick fish which may not be able to tolerate a “full” treatment.

It may be dosed by adding 0.5 ml per 10 gallons of water. This lower dose may also be appropriate for ponds in some situations; however, the use of formalin in ponds is often discouraged for several reasons, which are discussed below. Any time formalin is applied, vigorous aeration must be provided. When using formalin as a prolonged bath in a flow-through tank system, we recommend that the water flow be turned off for at least 12 hours (and up to 24 hours) to ensure a sufficient contact time with the parasite of concern.

Ensure that water quality (e.g., ammonia, nitrite, pH, dissolved oxygen) is optimal before stopping water flow and beginning the treatment. Research has shown that formalin at the above dosage rates does not have a significant effect on nitrifying bacteria in the biofilter of a recirculating system.

  1. However, higher dosage rates appear to have a negative impact on nitrite-oxidizing bacteria (the bacteria that convert nitrite to nitrate), which could lead to increases in nitrite concentration.
  2. Formalin will interfere with some water quality testing chemicals, most notably Nessler’s reagent, the chemical used in many ammonia test kits.

A salicylate-based test kit should be used to test ammonia whenever formalin is present in the water. In some situations, a high dosage rate of formalin up to 250 mg/l (1 ml formalin per gallon of water) may be applied as a short-term bath of 30–60 minutes.

  1. This treatment method is most appropriate in systems which have constant high water flow-rate, such as salmonid raceways, or where an individual or small number of fish will be moved into a new system with formalin-free water immediately after treatment.
  2. Tolerance of fish to this high dose rate and potential movement of pathogens if fish are moved to a new system must be considered.

Fish exhibiting signs of distress (e.g., darting, gasping, or trying to jump out of the water) during a chemical treatment should immediately be moved into clean (untreated) water.

Can formalin be used to preserve fish?

Formalin (formaldehyde in water) is a common adulterant in fish. Traders and suppliers use it to extend the storage life of fresh or chilled fish and artificially improve the sensory attributes.

How long does formalin stay active in water?

Supranee Chinabut, Chalor Limsuwan and Maliwan Sangjan ABSTRACT The toxicity of formalin at the concentrations of 25, 50 and 75 ppm to Aeromonas hydrophila was studied. A formalin concentration of 25 ppm killed 99.90 percent of bacteria within 24 hours.

Concentrations at 50 and 75 ppm killed 100 percent of A. hydrophila within 24 and 12 hours, respectively. Degradation of formalin was studied at the water temperature 27–30°C in aerated and non-aerated aquaria with and without fish. In the aerated aquaria with fish, formalin degraded more rapidly than in other groups.

At 25, 50 and 75 ppm, formalin completely degraded within 36, 48 and 54 hours in aerated group. In the non-aerated group, it did not degrade until after 36, 54 and 60 hours, respectively. In fiberglass containers, the degradation of formalin at the concentrations of 25 and 50 ppm was similar in the early hours (P 0.05).

  • Formalin at 25 and 50 ppm significantly reduced the amount of plankton and dissolved oxygen during the exposure time of 48 hours.
  • The results also showed that the total ammonia concentration slightly decreased in the early hours.
  • Other water quality parameters did not change.
  • INTRODUCTION The fish farming industry in Thailand is an expanding sector of the economy.

However, it has been recently beset by a number of major diseases which has increased the level of medication used by farmers. Apart from antibiotics used in the feed, the main medication taken in the form of external treatment via the water is formalin.

This compound is very active against external parasites which are themselves the cause of infections, or predispose the fish to other infections (Davis et al,, 1973; Goodman & Alfred 1975; Kabata 1985). It is likely that as fish culture becomes more intensive, formalin will be more widely used. Although some information on the toxicity and efficacy of formalin is available in developing countries, there is no published information on the degradation of formalin or its toxic effect on bacteria Aeromonas hydrophila which is the most important fish pathogen.

In order to use formalin effectively, the degradation rate must be known so that the chemical can be added into the water at the proper time. Thus, the specific objectives of the study are as follows:

To determine the toxicity of formalin to Aeromonas hydrophila, To determine the degradation rate of formalin in aquaria and fiberglass tanks. To determine the effect of formalin on water quality in fiberglass tank with high density of planktons. To determine the toxicity of formalin to planktons.

MATERIALS AND METHODS The experiments were conducted in the National Inland Fisheries Institute laboratory in Bangkok.1. The toxicity of formalin to Aeromonas hydrophila, A virulent strain of A. hydrophila isolated from a diseased walking catfish ( Clarias batrachus L.) was maintained on nutrient agar (NA) at 4°C.

  1. Then it was cultured on Brain Heart Infusion slant (BHI) and incubated for 18–20 hours at 30°C before the experiment started.
  2. Bacterial cultures were identified by the method described in Bergey’s manual (Buchanan & Gibbons 1974).
  3. The bacterial suspension was made by adding isotonic saline into the culture tubes until the optical density (O.D.) was 0.5 at 540 nm.

on a Baush & Lomb Spectronic 20 spectrophotometer. One ml of this bacterial suspension was added into each of the eight flasks which contained 78 ml. isotonic saline. One ml of formalin stock solution at the concentrations of 2000, 4000 and 6000 ppm were added into each flask so that the concentrations of formalin were 25, 50 and 75 ppm, respectively.

Distilled water was added to the control flasks. These experimental flasks were incubated at 30°C. Culture samples were withdrawn at 15 min, 1, 2, 3, 6, 12 and 24 hours and the viable counts were determined by the drop method (Collin & Lyne 1976). The effect of formalin on growth of A. hydrophila was compared with the untreated control.2.

The degradation rate of formalin in aquaria and fiberglass tanks. Aquaria Experiment Test fish and acclimation Common carps ( Cyprinus carpio Linn) weighing an average of 0.25– 0.4 gm were acclimated in aquaria (45×100×45 cm 3 ) containing well water and fed with pellet feed twice a day during the acclimation period.

  • Fish were not fed during the 24 hours before the test or during the test period.
  • Experimental design for aquaria There were four sets of experimental aquaria, aerated and nonaerated aquaria with and without fish present in the containers.
  • Two replications for each treatment were conducted.
  • Experiment with fish Twenty acclimated common carps were moved into each aquarium containing 100 litres of well water.

Formalin was added into the experimental aquaria at the concentrations of 25, 50 and 75 ppm. No formalin was added into the non-treated controlled aquaria. Formalin residue was measured following the method of AOAC (1984) at 0, 6, 12, 24, 36, 48, 54, 60, 72 hours after treatment.

Data were analyzed statistically by analysis of variance and Duncan’s new multiple range test (Steel & Torried 1986). Experiment without fish The experiment was conducted using the same methods as in the experiment with fish, described above. Experimental design for fiberglass tanks Five pairs of Nile tilapia ( Oreochromis niloticus, Linn) broodstock were moved into the fiberglass tanks which contained 1500 litres of green water from the fish pond.

The turbidity value was 30 cm. Fish were fed once a day with floating pellet at the rate of 3 percent of body weight through the experimental period. Formalin at the concentrations 25 and 50 ppm was added into the experimental tanks with two replications for each treatment.

  1. Formalin residue was measured following the method of AOAC (1984) at 0, 6, 30, 54 and 78 hours after treatment.3.
  2. To determine the effect of formalin on water quality in fiberglass tank with high density of planktons.
  3. Fiberglass tanks were filled with 1500 litres of green water from fish pond.
  4. The turbidity of water was 30 cm.

Five pairs of Nile tilapia were stocked per tank. Fish were fed once a day at 3 percent of body weight during the test period. Therapeutic levels of formalin at concentrations of 25 and 50 ppm were applied to each tank with two replications, and to untreated controls.

Analysis of water quality parameters, pH, temperature, dissolved oxygen, hardness, alkalinity and free carbon dioxide, of the experimental water was conducted according to the procedures outlined in Standard Methods for the Examination of Water and Wastewater (APHA) 1981). Total ammonia was determined by the method of Koroleff (Grasshoff 1976).

Turbidity values were determined at 0, 6, 30, 54 and 78 hours after treatment. Data was analysed using analysis of variance and Duncan’s new multiple range test (Steel & Torri 1986).4. To determine the toxicity of formalin to planktons. The experiment was conducted using the same qualities and measures in the third one.

  1. Plankton samples were collected at 0, 6, 30, 54 and 78 hours.
  2. The post-treatment plankton specimens preserved in 5 percent formalin were counted and identified.
  3. Statistical comparison was made among the treatments.
  4. RESULTS AND DISCUSSION Toxicity of formalin to Aeromonas hydrophila Formalin at a concentration of 25, 50 and 75 ppm each significantly killed Aeromonas hydrophila colonies more than 90 percent within one hour, with the degree of efficacy apparently being dose-dependent.

Table 1 and Fig.1 show that a formalin concentration of 25 ppm killed 99.90 percent of the bacteria A. hydrophila within 24 hours while formalin at the concentration of 50 and 75 ppm completely destroyed bacterial colonies within 24 and 12 hours, respectively.

Formalin Concentration (ppm.) Bacterial count
15 min 1 2 3 6 12 24
5.80×10 6 5.67×10 6 5.35×10 6 4.95×10 6 4.50×10 6 3.75×10 6 2.85×10 6
(6.76) (6.75) (6.73) (6.70) (6.65) (6.57) (6.45)
25 4.60×10 6 4.75×10 6 3.05×10 5 1.70×10 5 5.25×10 4 9.55×10 3 2.90×10 3
(6.66) (5.68) (5.49) (5.19) (4.72) (3.98) (3.46)
50 4.45×10 6 4.25×10 5 1.68×10 5 3.20×10 4 5.05×10 3 6.15×10 2
(6.65) (5.63) (5.23) (4.51) (3.71) (2.79)
75 4.20×10 6 3.15×10 5 1.07×10 5 1.35×10 4 2.30×10 3 1.38×10 2
(6.62) (5.49) (5.03) (4.12) (3.36) (2.14)

Remark Number in the parenthesis are logarithmic values of the mean

▟ CONTROL + 25 ppm. ◊ 50 ppm. 75ppm;

Fig.1. Toxicity of formalin to A. hydrophila The results of these experiments support the findings of earlier investigators, Braswell and his colleagues (1970) who reported that 15–25 ppm formalin applied to nutrient broth with bacteria significantly inhibited the growth rate of the bacteria.

  • The relatively large organic load of a fish pond might decrease the effective concentrations of formalin in the ponds and simultaneously provide a greater nutrient base for bacterial and protozoan repopulation.
  • Formalin concentration of 25 to 50 ppm provides effective reduction of ciliates protozoa (Bell et al,, 1987) and bacterial colonies in water and does not cause much damage to fish tissues.

This 25–50 ppm range of formalin concentration is the representative mode of formalin treatment. Degradation rate of formalin in aquaria and fiberglass tanks. Degradation rate of formalin in aquaria and fiberglass tanks was correlated to the time and test conditions. Fig.2. Percent degradation of formalin at various concentrations in aquaria with fish. (A) = aerated (NA) = non-aerated Fig.3. Percent degradation of formalin at various concentrations in aquaria without fish. (A) = aerated (NA) = non-aerated Formalin at 25 ppm was completely removed from the water within 36 hours in both aerated and non-aerated conditions. At the 75 ppm degradation rate was slower than at 25 and 50 ppm in both test conditions (Tables 2, 3, 4, 5, 6).

  1. In fiberglass tanks, the degradation rate of formalin was slightly slower than in aquaria (Table 7).
  2. The degradation of formalin at the concentrations of 25 to 50 ppm was similar in the early hours (P 0.05).
  3. In aerated condition, residual formalin at 24 hours after treatment from the starting concentrations of 25, 50 and 75 ppm were 1.31, 21.20 and 36.95 ppm, respectively.

Over the same time, 6.69, 30.44 and 43.69 ppm of residual formalin were detected in the non-aerated group which had the started concentration of 25, 50 and 75 ppm, respectively. This suggests that retreatment of formalin in the aquaria at the concentration of 25 ppm may be repeated at 24 hours after the first treatment.

  • Residual formalin from the fiberglass tanks containing green water at 24 hours after treatment from the starting concentrations of 25 and 50 ppm were 10.82 and 29.86 ppm, respectively.
  • Thus, 24 hours after the first treatment, 15 and 20 ppm of formalin can be added to maintain the concentration of 25 and 50 ppm.

Effect of formalin on water quality Dissolved oxygen was significantly reduced after application of formalin especially in the non-aerated aquaria (Fig.4, 5, 6, 7, 8). A similar conclusion was reported by Chalor (1985); Temdoung et al,, (1987) and Vinich et al,, (1987). Other water quality parameters were unchanged except pH and ammonia which slightly decreased after formalin application. The formic acid and the methylenetetramine compound produced from ammonia and formalin reaction (Tongchai 1982) may cause low pH and low ammonia in the water, respectively.

  • An indefinite treatment with formalin in green water pond may cause oxygen deficiency which is dangerous to some aquatic animals that cannot tolerate low oxygen condition.
  • Thus, aeration or water change is recommended in the formalin treatment ponds.
  • The toxicity of formalin to planktons The reduction in plankton counts in fiberglass tanks was correlated to the time and test conditions (Table 8, 9, 10).

Formalin at a concentration of 50 ppm reduced the number of plankton more significantly than 25 ppm and 0 ppm (control). Table II shows that 50 ppm formalin increases the turbidity value of the water more than 25 ppm does. RECOMMENDATIONS The findings of this study show that 25–50 ppm appears to be the appropriate formalin dose for pond treatment to reduce the levels of external fungi, protozoans, ectoparasitic monogeneans, crustaceans and bacteria A.

hydrophila, Oxygen deficiency may occur in the green water ponds after treatment with formalin, so that formalin treatment should be done in the early morning of a sunny day when photosynthesis of phytoplankton will increase oxygen content. REFERENCES Allison, R.1957. Some new results in the treatment of ponds to control some external parasites of fish.

Prog. Fish-Cult.19(2): 58–63. AOAC.1984. Official Methods of Analysis.14th ed., William Byrd Press, Inc., Virginia.1421 p. APHA, AWWA and WPCA.1981. Standard Methods for the Examination of Water and Wastewater.15th ed., American Public Health Association, Washington, D.C.1134 p.

  • Bell, T.A., C.S.
  • Arume and D.V.
  • Lightner.1987.
  • Effecacy of formalin reducing the levels of peritrichous ciliates on cultured marine shrimp.J. Fish.
  • Dis.10: 45–51.
  • Buchanan, R.E. and N.E.
  • Gibbons.1974.
  • Bergey’s Manual of Determinative Bacteriology.8th ed., The Williams and Wilkins co., Baltimore.1268 p.
  • Chalor, L.1985.

Fish Diseases. Fisheries Faculties, Kasetsart University, Bangkok.227 p. (In Thai). Collins, C.H. and P.M. Lyne.1976. Microbiological methods 4th ed., Butterworths. (Publishers) Inc., London.521 p. Davis, B.D., D. Renato, N.E. Herman, S.G. Harold and W. Barry.1973.

  1. Microbiology 2nd ed., Harper & Row Publisher, New York.1562 p.
  2. Goodman, L.S. and G.
  3. Alfred.1975.
  4. The Pharmacological Basis of Therapeuties.5th ed., Macmillan Publishing Co., Inc, New York.1704 p.
  5. Grasshoff, H.1976.
  6. Methods of Seawater Analysis.
  7. Verlag Chemic, New York.317 p.
  8. Abata, Z.1985.
  9. Parasites and Disease of Fish Culture in the Tropics.

Taylor & Francis, London.318 p. Roberts, R.J. and C.J. Shepherd.1979. Handbook of Trout and Salmon Disease. The Whitefriars Press Ltd., London.172 p. Steel, R.G.D. and J.H. Torrie.1986. Principles and Procedures in Statistics. McGraw-Hill, New York.633 p. Temdoung Pungkachonboon, C.

Limsuwan and S. Chinabut.1987. Toxicity of formalin to common carp, Cyprinus carpio L. In: Proceeding of the 25th Kasetsart University Conference, Fisheries Section, Kasetsart University, Bangkok.13–22 p. (In Thai). Vinich Tunsakul, Tongsuk Sai-Lee and Akaluck Sae-Loaw.1987. Acute toxicity and treatment effect of formalin to early larvae prawn, Macrobrachium rosenbergii (De Man).

In: Proceeding of the 25th Kasetsart University Conference Fisheries Section, Kasetsart University, Bangkok, 23–33 p. (In Thai). Wellborn, T.L.1979. Control and therapy, pp 61–62. In: J.A. Plumb (ed.). Principle Disease of Farm-Raised Catfish. Sothern cooperative Series No.225.

Formalin Concentration (ppm) percent degradation of formalin *
6 12 24 36 48 54 60 hours
25 (A) 27.42 a 54.38 a 95.41 a 100 a 100 100 100
25 (NA) 16.30 b 31.43 b 76.29 b 100 a 100 100 100
50 (A) 19.10 a 27.58 a 61.74 a 84.38 a 100 a 100 a 100
50 (NA) 7.58 b 18.86 b 44.82 b 67.72 b 94.26 b 100 a 100
75 (A) 10.98 a 25.37 a 54.48 a 74.83 a 91.17 a 100 a 100 a
75 (NA) 6.82 b 14.82 b 45.55 b 71.64 b 85.77 b 96.27 b 100 a

Remark (A) = aerated (NA) = non-aerated * Values within a column of the same concentration and a different letter are significantly different. Table 3, The percentage of formalin degradation rate at various time scales after treatment in aquaria without fish.

Formalin Concentration (ppm.) percent degradation of formalin *
6 12 24 36 48 54 60 hours
25 (A) 16.09 a 28.57 a 72.54 a 98.17 a 100 a 100 100
25 (NA) 8.49 b 18.13 b 59.73 b 86.84 b 100 a 100 100
50 (A) 11.83 a 21.13 a 53.66 a 75.74 a 95.13 a 100 a 100
50 (NA) 6.35 b 13.59 b 39.41 b 60.35 b 87.06 b 100 a 100
75 (A) 9.12 a 17.25 a 38.34 a 65.83 a 81.88 a 95.46 a 100 a
75 (NA) 4.21 b 10.41 b 27.25 b 51.85 b 74.56 b 89.03 b 100 a

Remark (A) = aerated (NA) = non-aerated * Values within a column of the same concentration and with different letters are significantly different. Table 4, The concentration of formalin residual at various time scales after treatment in the aquaria without fish.

Formalin Treatment Formalin residual (ppm)
Concentration (ppm) 6 12 24 36 48 54 60 hours
25 28.12 23.60 20.09 7.72 0.52
50 55.78 49.18 44.06 25.85 13.53 2.71
75 82.43 74.94 68.21 50.84 28.18 14.94 3.71
25 27.23 24.92 22.30 10.97 3.59
50 56.52 52.94 48.84 35.26 22.40 7.33
75 80.69 77.29 72.28 58.71 38.85 20.55 8.22

Remark Treatment 1 = aerated Treatment 2 = non-aerated Table 5, Percent degradation of formalin at various time scales after formalin treatment with aeration.

Formalin Concentration (ppm) percent degradation of formalin *
6 12 24 36 48 54 60 hours
25 (F) 27.42 a 54.38 a 95.41 a 100 a 100 a 100 100
25 (NF) 16.09 b 28.57 b 72.54 b 98.17 a 100 a 100 100
50 (F) 19.10 a 27.58 a 61.74 a 84.38 a 100 a 100 a 100
50 (NF) 11.83 b 21.13 b 53.66 b 75.74 b 95.13 b 100 a 100
75 (F) 10.98 a 25.37 a 54.48 a 74.83 a 91.17 a 100 a 100 a
75 (NF) 9.12 a 17.25 b 38.34 b 65.83 b 81.98 b 95.46 b 100 a

Remark (F) = with fish (NF) = without fish * Values within a column of the same concentration and with a different letters are significantly different. Table 6. Percent degradation of formalin at various time scales after formalin treatment in non-aerated experiment

Formalin Concentration (ppm) percent degradation of formalin *
6 12 24 36 48 54 60 hours
25 (F) 16.30 a 31.43 a 76.29 a 100 a 100 a 100 100
25 (NF) 8.49 b 18.13 b 59.73 b 86.84 b 100 a 100 100
50 (F) 7.58 a 18.56 a 44.82 a 67.72 a 94.26 a 100 a 100
50 (NF) 6.35 a 13.59 b 39.41 b 60.35 b 87.06 b 100 a 100
75 (F) 6.82 a 14.82 a 45.55 a 71.64 a 85.77 a 96.27 a 100 a
75 (NF) 4.21 a 10.41 b 27.25 b 51.85 b 74.56 b 89.83 b 100 a

Remark (F) = with fish (NF) = without fish * Values within a column of the same concentration and with a different letter are significantly different. Table 7, Percent degradation of formalin at various time scales after formalin treatment in fiberglass tank with fish.

Formalin Concentration (ppm) percent degradation of formalin
6 12 24 36 48 54 60 hours
25 19.25 a 31.60 b 58.02 c 79.53 d 100 h 100 h 100 h
50 14.35 a 25.25 b 42.74 c 59.85 e 81.02 f 92.83 g 100 h

Remark * Values within a column of the same concentration but with a different letter are significantly different. Time (hours) Fig.4 Water quality aerated aquaria with fish after formalin treatment Time (hours) Fig.5 Water quality in non-aerated with fish after formalin treatment Time (hours) Fig.6 Water quality in aerated aquaria without fish after formalin treatment Time (hours) Fig.7 Water quality in non-aerated aquaria without fish after formalin treatment Time (hours) Fig.8 Water quality in fiber-glass tank after formalin treatment Table 8, Numbers of planktons in untreated control fiberglass tank experiment at various time scales.

Plankton No. of plankton (cell/liter)
Before treatment treatment
6 30 54 78 hours
Coelastrum 1,521 1,459 1,253 894 835 328
Synedra 1,357 1,236 1,020 1,003 668 365
Dictyosphaerium 1,303 1,251 859 798 572 201
Anabaenopsis 981 902 852 821 547 281
Oocystis 857 841 784 674 434 206
Desmid 509 497 256 208 137 91
Scenedesmus 49 48 39 33 16 18
Pediastrum 79 27 22 21 16 16
Chroococcus 48 42 38 34 32
Staurastrum 18 17 12 11 11 9
Oedogonium 9 6 6 4 3 3
Keratella 12 9 7 7 3 2
Anabaena 3 2
Brachionus 90 79 60 25 18 18
Filinia 8 5 3 3 2
Nauplius 12 12 11 9 3
Copepod 4 3 3 2
Unidentified-green algae 5 5 3 3 2
Unidentified blue-green algae 139 139 138 123 73 59
Total 7,004 6,580 5,366 4,673 3,372 1,597

Table 9, Numbers of planktons in fiberglass tank experiment before and after treated with formalin at the concentration of 25 ppm.

Type of Plankton No. of plankton (cell/liter)
Before treatment Treatment
6 30 54 78 hours
Coelastrum 1,530 1,304 1,100 884 502 57
Synedra 1,415 1,228 810 774 522 274
Dictyosphaerium 1,282 1,030 663 584 256 57
Anabaenopsis 976 886 818 704 223 45
Oocystis 866 820 644 573 374 51
Desmid 534 491 264 241 112 58
Scenedesmus 81 74 53 38 18 7
Pediastrum 68 40 30 21 8 5
Chroococcus 68 44 43 42 39 2
Staurastrum 18 16 14 12 10 5
Oedogonium 26 16 10 8 4 2
Keratella 16 14 14 12 4 2
Anabaena 6 4 4 2
Brachionus 78 64 24 21 10 3
Filinia 10 8 6 6 2 1
Nauplius 17 4 4 2 1
Copepod 12 8 6 3 2 2
Unidentified-green algae 8 8 7 6 6
Unidentified blue-green algae 188 175 133 99 76 33
Total 7,199 6,234 4,647 4,032 2,169 604

Table 10, Numbers of plankton in fiberglass tank experiment before and after formalin treatment at the concentration of 50 ppm.

Type of plankton No. of plankton (cell/liter)
Before treatment Treatment
6 30 54 78 hours
Coelastrum 1,362 1,180 1,020 418 214 51
Synedra 1,226 965 791 422 404 229
Dictyosphaerium 1,248 779 531 194 118 28
Anabaenopsis 994 809 757 340 108 43
Oocystis 858 642 502 243 193 44
Desmid 519 331 122 54 43 27
Scenedesmus 76 69 41 24 15 5
Pediastrum 58 51 38 38 8 8
Chroococcus 48 22 17 14 13 2
Staurastrum 23 16 8 5 3 3
Oedogonium 10 8 6 4 2 2
Keratella 16 14 12 2 2 1
Anabaena 8 2 2 1 1
Brachionus 50 48 37 14 8 3
Filinia 15 5 4 2 1
Nauplius 6 3 2 1
Copepod 6 6 4 2 2 1
Unidentified-green algae 11 8 6 2 1
Unidentified blue-green algae 163 142 73 56 54 17
Total 6,697 5,100 3,973 1,835 1,190 465
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Table 11, Water turbidity in fiberglass tank experiment before and after formalin treatment at the concentration of 25 and 50 ppm.

Formalin Concentration (ppm.) Turbidity (cm.)
Before treatment Treatment
6 30 54 78 hours
Control 33.0 33.0 34.0 33.0 28.4 25
25 34.0 34.5 36.5 43.0 49.3 47
50 34.5 35.0 40.5 53.5 54.5 58.5

What is formalin solution used for?

Uses of Formelene –

Formalin solution can be used as a disinfectant because it destroys most bacteria and fungi (including their spores). Toxins and viruses are inactivated using it as an ingredient in vaccine production. In personal care products like cosmetics, formaldehyde releasers are used as biocides. They are known to cause allergic contact dermatitis in some sensitized individuals, despite being present at levels that are not usually considered dangerous. It’s used to create resins like urea formaldehyde and phenol-formaldehyde, which are used in foundries to make cores and molds. It’s used as a disinfectant, fungicide, fumigant, and preservative in agriculture and medicine. A broad range of products, especially building materials, contain formaldehyde.

Can you mix formalin and water?

To make a histological fixative from this we need a 10% solution** of this stock formalin i.e.1 part of the stock formalin with 9 parts water, preferably distilled. This makes an unbuffered formalin solution, which will have a pH of 3-4.

Does formalin react with water?

Abstract – Formaldehyde is a hydrogen-containing chemical that can react with water, and thus, formaldehyde–water has emerged as an attractive hydrogen storage system. However, formaldehyde is also a carcinogenic airborne pollutant released from paint, coatings and furniture.

  • In this work, we innovatively propose a method for Ru( p -cymene) immobilization and prepare several novel ruthenium organic crystalline particles for formaldehyde decomposition.
  • The ruthenium organic crystalline particles are bifunctional catalysts.
  • On one hand, they exhibit high activity for hydrogen production from formaldehyde–water decomposition in water.

The TOF reaches up to 2420 h −1 at 90 °C when Ru-DAPM 2 is employed. On the other hand, ruthenium organic crystalline particles also have high activity for gaseous formaldehyde decomposition in air. Unlike the reported photocatalytic or thermocatalytic oxidation methods for gaseous formaldehyde removal, the ruthenium organic crystalline particles provided two reaction paths for formaldehyde decomposition: formaldehyde–water decomposition or the catalytic oxidation of formaldehyde.

  • Formaldehyde mainly decomposes through water-induced reactions in moist air, and Ru-DAPM 2 exhibits 98% conversion for formaldehyde decomposition at 90 °C.
  • We propose and prove the reaction process and mechanism for formaldehyde decomposition in air, and find that Ru-DAPM 2 first absorbs water vapor in air.

The gaseous formaldehyde then dissolves in water, is converted into methanediol, undergoes the formaldehyde–water shift reaction, and produces hydrogen and formic acid. Formic acid is an essential intermediate during the reaction, and could further decompose into hydrogen and carbon dioxide.

Can bacteria survive in formalin?

Presence of Contagious Bacterial Flora in Formalin-Fixed Cadavers: A Potential Health Hazard to Medical Professionals Cadaveric dissection is the most important learning tool in anatomy. Although many new modalities are coming up for learning anatomy, cadaveric dissection outstands all of these as it helps students to visualize the human body and remains the most realistic way of learning anatomy.

  1. The cadavers are preserved using formalin, a potent disinfectant.
  2. Even after embalming in 5-10% formalin, the cadaver might still be infectious while using it in the dissection hall (anatomy department).
  3. Numerous bacterial species and infectious pathogens might still be seen despite using fixative agents.

Several disease-causing agents may remain viable. Earlier reports suggest that there are cases where students and the working staff got infected by HIV, viral hepatitis, tuberculosis, and prion diseases. The main objective of this study is to determine if bacterial species could be recovered from cadavers that are formalin-fixed.

  • Specific regions in the body such as the axilla, perineum, finger clefts, and oral and nasal cavities were chosen for microbiological examination to detect bacterial species.
  • The presence of skin folds in these regions makes them potential sites for the growth of bacteria.
  • Formalin-fixed cadavers can still act as regions for the growth of viable bacteria that can be pathogenic and affect the health of students and anatomists handling them.

Proper care should be taken regarding this because students and anatomists working with these cadavers may get exposed to pathogenic organisms which may become harmful or sometimes life-threatening. Some precautions for proper dissemination of cadavers should be taken to provide a complete, safe, and healthy ambiance in the dissection hall.

  • Eywords: formalin, disinfection, bacteria, dissection halls, cadavers Cadavers have been one of the best tools to learn human anatomy for centuries.
  • They help students learn about muscles, bones, vessels, and every minute structure of the human body in a very detailed manner,
  • This gives them more accurate knowledge about the size and location of every organ inside the body.

Learning with the help of a cadaver teaches teamwork as a group of students works together at dissection hall tables helping out each other, Cadaveric dissection makes anatomy more interesting and creates enthusiasm among students. The most common source of cadavers in dissection halls are unclaimed bodies or donated bodies of individuals.

A few anatomists also import cadavers from other countries to conduct a variety of studies, The cadavers have to be checked for the presence of microorganisms before handling them to students in the dissection hall. The main reason for embalming is to inhibit the growth of microorganisms and preserve the tissue for longer studies,

The commonly used embalming fluids include formalin, ethanol, and phenol, which are effective against infective agents such as bacteria, fungi, and spores. Formalin is an aqueous solution of formaldehyde that acts by the formation of covalent bonds, thus inhibiting the growth of various bacteria and fungi.

Few studies indicate formalin being ineffective in removing all microorganisms from the surface of cadavers, while few show differences in the usage of formalin percentage that would affect the growth of microorganisms completely, There are numerous viable organisms which include bacteria ( Staphylococci, Streptococci, etc.) fungi ( Penicillium, Aspergillus, etc.), and viruses present on formalin-fixed cadavers that can contaminate the dissection halls and transmit various diseases to students if proper care is not taken.

They may transmit diseases such as HIV, hepatitis B and C, and tuberculosis to people handling cadavers in the dissection hall, Proper protective clothing, maintenance of hygiene, vaccination against diseases such as hepatitis and tuberculosis to all those who handle cadavers, and staying up to date on the most recent literature in the field would be of great help to ensure safety,

This study would give us adequate knowledge: (1) to check for the presence of bacteria on formalin-fixed cadavers and determine whether they are commensal or pathogenic; and (2) to prevent anatomists and students from getting exposed to pathogenic organisms while handling the cadavers. Common risk factors and their adverse effects Chemicals Formalin is the most commonly used chemical substance to preserve cadavers according to our regular practices.

This is found to show various adverse effects on the students and dissectors which include headache, nausea, dizziness, dryness of eyes and mucous membranes, overflow of tears, and a burning sensation in the eyes and throat. Long-term exposure to formalin may also lead to skin disorders and cancer,

Unhygienic PracticesUnprotected and improper clothing (laboratory coats), unclean hands, and instruments can act as carriers as various pathogens get lodged on them and can get administered into the body through any route,Inadequate Ventilation and Ineffective Laboratory Practices

Several students working together in closed rooms/halls during dissection with improper ventilation can increase the chances of transmission of airborne diseases. Laboratory practices such as improper disposal of the dissection hall waste such as tissues and skin, inadequate disinfection, and preservation techniques of the cadavers are major risk factors for various disease transmission,

  1. Specific body regions and the various bacterial species that are detected According to the study that was conducted, specific regions of the body like the axilla, oral, nasal, and perineal regions of the cadavers were examined to check for the presence of bacteria,
  2. The presence of skin folds in these regions makes them potential sites for the growth of bacteria.

The particular regions of the sample collection are shown in Figure, The identified bacterial species were further labeled as gram-positive or gram-negative species. The specific body region that they were identified from is mentioned in Table, Bacteria cultured from the surfaces of formalin-fixed cadavers.

Bacterial species Body region Common infections caused
Aerococcus viridans Axilla Bacteraemia, endocarditis, urinary traction infections
Cellulomonas Perineum Bacteremia and catheter-related infections
Gardenerella vaginalis Axilla Bacterial vaginosis
Gemella hemolysans Perineum Eye infections, meningitis
Staphylococcus auricularis Axilla Skin and soft-tissue infections
Staphylococcus epidermidis Axilla, oronasal, perineum Bloodstream infections
Staphylococcus haemolyticus Axilla, perineum Bacteremia, meningitis, endocarditis
Strepto coccus mitis Oronasal Dental and eye infections, infective endocarditis, bacteremia
Staphylococcus lugdunesis Oronasal Skin and soft-tissue infections, endocarditis
Corynebacterium striatum Perineum Bloodstream and catheter-related infections
Corynebacterium propinquum Perineum Infective endocarditis, respiratory tract infections
Kocuria varians Oronasal Urinary tract infections, peritonitis, bacteremia
Kocuria kristinae Oronasal, perineum Urinary tract infections
Gemella morbillorum Axilla, oronasal Infective endocarditis

A brief review of all identified bacterial species Aerococcus viridians These are gram-positive organisms that belong to the Aerococcus group and are considered contaminants that are seen in the hospital environment. Aerococcus was first described under a single species A.

  1. Viridans ; further, five more new species were identified, namely, A.
  2. Urinae, A.
  3. Sanguinicola, A.
  4. Christensenii, A.
  5. Urinaehominis, and A.
  6. Urinaeequi,
  7. It is used as a source of lactate oxidase commercially.
  8. This is a gram-positive coccus and has a morphology similar to that of viridans streptococci.
  9. It is a rare pathogen that usually causes urinary tract infections, endocarditis, bacteremia, arthritis, or meningitis.

Aerococcus shows fastidious growth and is usually confused with strains of Streptococcus and Staphylococcus, Aerococcus infections are usually treated by giving intravenous (IV) penicillin or ceftriaxone monotherapy, Cellulomonas It is a gram-positive, rod-shaped bacillus that has a special ability to degrade cellulose using specific enzymes such as endoglucanase and exoglucanase.

  1. They belong to the Actinobacteria group.
  2. This pathogen is also not very commonly seen, but it is recently noted to be one of the emerging pathogens for humans.
  3. Few cases have been reported where it caused endocarditis and osteomyelitis.
  4. Identification of cellulosomes on cadavers is unusual as it is usually found in soil and has cellulose activity.

Cellulomonas strains usually possess various genes for the production of cellulases and xylanases, Gardnerella vaginalis The organism was named Haemophilus vaginalis when it was first identified. It is a facultative anaerobe that shows gram-variable staining because of the presence of a thin cell wall.

These are non-motile and do not form spores. Gardnerella is normally found as a common organism of vaginal flora and contributes to maintaining balanced pH in the region. It causes bacterial vaginosis when it starts growing abnormally by involving other anaerobic bacteria and destroys the normal vaginal flora.

It produces a toxin called vaginosis that affects human cells specifically, Gemella hemolysins It is a gram-positive, facultative anaerobe. They resemble Neisseria species. Gemella species are usually found in the oral cavity and respiratory tract of human beings where there is the presence of mucous membranes.G.

  • Hemolysins species is usually isolated from nasopharyngeal swabs of people.
  • It is known to cause pulmonary exacerbations in patients with cystic fibrosis.
  • The presence of this organism in the saliva is related to the periodontal health of the oral cavity and prevents the growth of P.
  • Gingivalis,
  • Staphylococcus auricularis It is a gram-positive bacteria present in pairs or tetrads.

This is usually found in the head region where abundant sebaceous glands are found, especially in the exterior region of the ear (auditory canal). It is weakly hemolytic. It is commonly found on human skin, causing sepsis or any other infections, but is usually a rare coagulase-negative Staphylococci (CoNS).

  1. Skin and soft-tissue infections are usually seen,
  2. Staphylococcus epidermidis It is a gram-positive bacteria and one of the common microflora that infects human epithelium.
  3. This typically belongs to human skin flora.
  4. The bacteria are usually found in hidden regions of the body and can cause boils and infections.

It is an important opportunistic pathogen that gets easily attached to catheters and other medical devices to form biofilms. It is most commonly found in the axilla, head, and nostrils and belongs to the group of CoNS. These infections are usually treated with penicillin G, cephalosporins, etc.

Vancomycin is used as the drug of choice for methicillin-resistant organisms, Staphylococcus haemolyticus It is a gram-positive coccus and an opportunistic bacterial pathogen. It is usually found in the axilla, perineum, and inguinal regions and can cause bacteremia, meningitis, and skin or soft-tissue infections.

This also has extreme antibiotic resistance and can be transmitted through direct or indirect contact with an infected person.S. hemolyticus can be rapidly inactivated through dry Cu. These infections are also usually hospital-acquired due to the administration of medical devices.

The formation of biofilms makes the treatment of this bacteria more difficult. This organism also colonizes domestic animals, Staphylococcus lugdunensis It is a gram-positive bacteria that belongs to CoNS. They are usually hemolytic, appear in clusters, and have a sweet-smelling odor. It causes skin and soft-tissue infections such as cellulitis, cystic lesions, and peri-inguinal abscesses.

It also causes a severe form of endocarditis, osteomyelitis, endocarditis, arthritis, and septicemia. It should be looked into for all routine investigations in the laboratories because they are one of the main pathogens for skin and soft-tissue investigations, especially in the groin region leading to cellulitis and abscesses,

  • Streptococcus mitis S.
  • Mitis is gram-positive cocci that are spherical facultative anaerobe.
  • It is an alpha-hemolytic species of Streptococcus,
  • This bacteria was previously known as S.
  • Mitior and is usually found in the mouth, throat, nasopharynx, female genital tract, gastrointestinal tract, and skin.

It has less virulence and pathogenicity but may cause severe infections such as endocarditis and meningitis which are life-threatening. Nearly 20 strains of S. mitis have been isolated from body fluids and are found to be susceptible to optochin. These bacteria move through the bloodstream to specifically get attached,

Corynebacterium striatum This is a gram-positive, non-diptherial corynebacterium with a club-shaped morphology. They usually contaminate human skin and mucous membranes. They cause several infections including bacteremia, meningitis, pleuropneumonia, osteomyelitis, and uterine infections. It is one of the most emerging pathogens in various countries and is one of the most frequently isolated species of Corynebacterium,

These bacterial infections are usually treated with monotherapy or amoxicillin-clavulanic acid in mild conditions, and in severe conditions, daptomycin or linezolid is used. It is a multi-drug-resistant pathogen that usually colonizes the nasopharynx,

  • Corynebacterium propinquum It is a gram-positive aerobic bacteria that is normally a part of the oropharyngeal flora that is found in the skin and mucous membranes of the respiratory system.
  • It is primarily isolated from the respiratory tract.
  • It has been detected as a pathogen in infective endocarditis, prosthetic valve endocarditis, and nosocomial infections.

These infections are usually seen in immune-compromised patients or those with underlying lung diseases. This can also cause other respiratory infections such as chronic obstructive pulmonary disease, bronchiectasis, and pneumonia. This species shows resistance to vancomycin and daptomycin.

  1. Proper identification of the species and the extent to which it could be pathogenic should be known to provide specific treatment,
  2. Ocuria varians It is a gram-positive bacteria that resembles Staphylococcus and Micrococcus and is arranged in pairs or tetrads.
  3. They are either aerobic or facultative anaerobic that have very rigid cell walls.

It is generally found live on the skin and oral cavity. This bacteria is usually non-pathogenic but is related to specific infections such as urinary tract infections, cholecystitis, peritonitis, catheter-associated infections, brain abscess, and meningitis.

Kocuria causes infections in immunosuppressed patients related to the oropharynx and deep cervical lymph nodes. These organisms grow on simple media plates or blood agar. They do not have the hemolytic ability on blood agar. Kocuria shows resistance to lysostaphin and nitrofurantoin, Kocuria kristinae It is a gram-positive bacteria that is formerly known as Micrococcus kristinae,

This is normally found on human skin and oral mucosa and causes opportunistic infections. Recently, this organism was named Rothia kristinae after reclassifying the species. It is a facultative, non-motile anaerobe that forms pale cream to pale orange colonies on blood agar.R.

  • Kristinae is resistant to lysozyme.
  • This organism is not considered to be a primary pathogen but is seen in patients under hospitalization, catheter-related bacteremia, patients undergoing dialysis, pregnant females, or patients with chronic illness.
  • Treatment of this organism involves parenteral administration of vancomycin along with some other antibiotics,

Gemella morbillorum It is a gram-positive coccus that lives in a microaerophilic environment. It was formerly known as Streptococcus morbillorum, It rarely causes any diseases, although it is found in the oropharyngeal area and gastrointestinal tract.G.

  • Morbillorum predominantly causes endocarditis, endovascular infections, and a few invasive infections.
  • It is one of the most common bacteria found on teeth with cysts that do not get resolved even after repeated root canal treatments.
  • These bacterial strains are usually resistant to penicillin.G.
  • Morbillorum has also been seen in colorectal cancer.

It is one of the most common organisms to cause infective endocarditis in children. The most common cause of G. morbillorum is poor dental health or oral hygiene. It is a commensal of the upper respiratory tract of humans and gets into the bloodstream of patients to cause bacteremia,

Implications This study helps us to know the presence of bacterial flora in different regions of the body, the importance of standard infection control methods that have to be followed by students and anatomists in the dissection hall to avoid the spread of any infections, and the need to improvise the dissemination methods that are currently being followed.

Proper clothing (laboratory coat), covering hair (using cap masks), hand gloves, table sterilization, proper disposal of unwanted tissues and debris after dissection, post-dissection hand washing and usage of hand sanitizers, and vaccination may prevent the spread of infections to some extent.

  • The dissection hall should be maintained properly by proper cleaning with disinfectants such as phenol or formaldehyde.
  • Covering the cadaver properly after usage with a sheet and disposing of all the waste as per the guidelines of biomedical waste management should be made compulsory.
  • Hand washing limits the spread of the disease to a greater extent and should be well understood and followed by all the students and staff.

Proper vaccination against hepatitis B, tuberculosis, etc. to avoid their transmission in accidental injuries while handling the cadavers is crucial. As formalin also causes a lot of discomfort to students leading to watery eyes, dryness of mucous membranes, headache, nausea, and gastrointestinal disturbances, studies are ongoing to come up with better methods that can replace the usage of formalin or in alternate concentrations that can be completely effective for disinfection.

  1. It is required to look for an alternative to formalin as it is dangerous and hardens the tissues making dissection hard for students.
  2. Medical students, faculty members, and cadaver handlers who come in contact with cadavers in dissection halls are prone to get infected by these microorganisms.
  3. Formaldehyde fumes are harmful and cause skin and mucous membrane irritation.

Researchers are now trying to identify different embalming procedures with a wide variety of chemicals instead of formalin. Above all, formalin is still used as a preservative and disinfectant in dissection halls because of its low cost, effectiveness, and results.

Therefore, it is recommended to take proper precautions to avoid unnecessary exposure to chemicals and microorganisms until a proper alternative is identified. Numerous bacterial species have been identified in formalin-fixed cadavers that are given to medical students for dissection. For studying these bacteria, specific sites in the human body such as axillary, perineal, and oronasal regions were chosen as they are not usually contacted by people.

These regions usually have skin folds that can prevent them from direct contact with formalin. Identification of bacterial flora will help us understand various kinds of diseases that can get transmitted to people dealing with cadavers. This clearly tells us the need to follow proper precautions while handling cadavers in dissection halls.

  1. The content published in Cureus is the result of clinical experience and/or research by independent individuals or organizations.
  2. Cureus is not responsible for the scientific accuracy or reliability of data or conclusions published herein.
  3. All content published within Cureus is intended only for educational, research and reference purposes.

Additionally, articles published within Cureus should not be deemed a suitable substitute for the advice of a qualified health care professional. Do not disregard or avoid professional medical advice due to content published within Cureus. The authors have declared that no competing interests exist.1.

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Is formalin toxic to humans?

What immediate health effects can be caused by exposure to formaldehyde? – Formaldehyde can cause irritation of the eyes, nose, and throat, even at low levels for short periods. Longer exposure or higher doses can cause coughing or choking. Severe exposure can cause death from throat swelling or from chemical burns to the lungs.

Direct contact with the skin, eyes, or gastrointestinal tract can cause serious burns. Drinking as little as 30 mL (about 2 tablespoons) of formalin can cause death. Formate, a formaldehyde metabolite, can cause death or serious systemic effects. Generally, the more serious the exposure to formaldehyde, the more severe the symptoms.

Previously sensitized persons may develop a skin rash or breathing problems from very small exposures.

What is a good container for formalin?

Histoplex™ Histology Containers forms a seal that eliminates most formalin leakage and evaporation. The durable polypropylene container (5ml-1 liter) is translucent, flexible, resists cracking and splitting, and eliminates the problems associated with formalin evaporation.

How long is 10% formalin good for?

International Specimen Collection Recommendations and Submission Guidelines for Pathologic Evaluation of Unexplained Illness due to Possibly Infectious Etiology Determining the cause of an infectious disease can be challenging, particularly when specimens such as serum, blood, or cerebrospinal fluid (CSF), a collection of the fluid around the brain and spinal cord, are unavailable, or when traditional laboratory assays have been unsuccessful at identifying a specific agent.

Tissue-based diagnostics, utilizing specimens acquired through biopsy or autopsy provide extremely powerful and versatile methods of establishing an etiologic diagnosis. These samples, when submerged in 10% neutral buffered formalin (up to 14 days for biopsy or autopsy tissue, or up to 28 days for brain autopsy tissue), are almost always rendered non-infectious and can be transferred to 70% ethanol for long-term storage at room temperature until analysis by specific histopathologic, immunohistochemical or molecular assays.

For deceased patients, tissues obtained at autopsy are preferable; however, when an autopsy is impractical because of real or perceived biosafety risks, or because of cultural practices, limited tissue collection may still be possible and should be considered.

When an autopsy is performed representative portions from all major organs (lung, heart, liver, kidney, spleen, and brain), as well as any tissues or organs that show conspicuous lesions or are indicated by clinical findings, should be submitted for evaluation. When a complete autopsy of a deceased patient is not possible a full-thickness biopsy should be collected from any conspicuous skin lesion (petechiae, eschar, or purpura). Core or wedge biopsy specimens of major organs (particularly liver), obtained by using a viscertome (a device used to get a liver tissue sample) or by limited autopsy, are also acceptable.

Biopsy specimens of skin, bone marrow, or other tissues, obtained from living patients, are also acceptable for evaluation. All tissue specimens should be fixed in 10% neutral buffered formalin for a minimum of 3 days (72 hours) and up to 2 weeks (14 days) for biopsy or autopsy tissue, or up to 4 weeks (28 days) for brain autopsy tissue.

Biopsy tissues should be paraffin-embedded prior to submission to IDPB. If not paraffin-embedded, after a minimum of 3 days (72 hours) of formalin-fixation, autopsy tissues can be transferred to 70% ethanol for long term storage and shipping at room temperature. Each specimen MUST be labeled clearly with a unique patient identifier that includes the patient’s name,

When available, a brief clinical history, including any pertinent laboratory results, should also be included.

Does 10% formalin go bad?

Variables that influence formalin fixation – Post-Mortem or Post-Surgical Interval. Autolysis begins immediately following disruption of blood flow and rapid fixation is essential. Because autolysis can influence the cellular morphology and staining, the time interval between death and fixation should be minimized and controlled whenever possible. If there is a necessary delay in fixation, the tissue should be immersed in cold phosphate-buffered saline (PBS). Tissues should not be allowed to dry before (or after) fixation. For certain studies, vascular perfusion with fixative may be recommended. Fixative Composition. Formaldehyde fixation is usually optimal near physiological pH and ionic strength. Unfortunately, formalin is not stable and will gradually acidify and form more complex polymers, with adverse effects on fixation. This is minimized by appropriate buffering and inclusion of small amounts of methanol. Nevertheless, formalin has a finite shelf-life. Commercial preparations in specimen containers are strongly recommended, and should not be used beyond the expiration date, usually 2 years. Fixative Volume. Tissues should be immersed in a >20-fold volume excess of fixative. Large amounts of blood or protein-rich fluid can decrease the effective fixative concentration. If necessary, this problem can be minimized by brief washing of the tissue with PBS prior to placement in the fixative. If the fixative becomes obviously cloudy or bloody the fixative should be changed. Fixation Time. Fixation with 10% NBF at room temperature usually provides excellent morphological detail. For most animal tissues, a minimum of 24 hours at room temperature is recommended. However, some bloody or fatty tissues, and certain fetal tissues, could require significantly longer fixation, e.g., up to 48- 72 hours. If sectioning of a fixed tissue reveals internal areas with normal coloration, the fixation time should be extended. Small clinical biopsies are sometimes fixed for only 6 hours, but 12 or more hours will often provide a better result. As suggested above, under-fixation is more of a problem than over-fixation for most histochemical procedures. Under-fixation can be associated with artifacts secondary to alcohol exposure during tissue processing, or tissue damage during decalcification procedures. Over-fixation can impair staining, and complicate sectioning, but can more often be compensated for by the histotechnologist. Fixation Temperature. Fixation time and temperature are related variables. An increase in temperature can increase the rate of fixation but can also increase the rate of autolysis. Primary fixation in the cold can slow autolysis, but also slows the process of fixation, and premature cooling of a specimen in fixative can lead to inadequate fixation. Tissue Thickness. Formalin fixation is dependent on diffusion of the fixative into the tissue. The thicker the tissue, the more time is required to obtain complete penetration and cross-linking of tissue proteins. If the specimen is too thick, the center of the tissue can suffer from autolysis or incomplete fixation. For this reason, tissues should be no thicker than 3 mm. Careful slicing of tissues and solid organs prior to transfer to fixative can greatly influence the efficiency of fixation by increasing exposed surface area and decreasing total thickness. This is particularly important for fatty, bloody, or organs surfaced by a capsule or membrane. When slicing tissues, it is important to use a sharp, clean blade to minimize compression artifacts. If forceps are used to stabilize the tissue, care must be taken to avoid applying excessive pressure. For very soft tissues, it is sometimes helpful to stiffen the tissue by brief fixation prior to cutting of thinner sections and additional fixation. Tubular organs can be cross-sectioned at small intervals to enhance exposure of the lumen to the fixative; any luminal contents should also be removed. Post-Fixation Storage. Non-coagulant fixatives such as formalin continue to cross-link proteins as long as they are in contact with the tissue. If there is to be a delay in processing after complete fixation (usually 24 hours or more), tissue can be stored for up to 3 days in the cold in 70% ethanol. However, it is essential that the tissue is completely fixed prior to transfer to the alcohol. Premature transfer of incompletely fixed tissue to alcohol precludes normal tissue processing has been reported to permit time-dependent reversal of cross-linking.

How long does it take for formalin to penetrate tissue?

Formalin penetrates tissues slowly ( approx.1mm per hour ) 1,9 so specimens need to be opened, incised or sliced and left to fix for an adequate period of time prior to processing.

Why formalin is banned?

2. Prohibition of formaldehyde: The dangers of formaldehyde : – Formaldehyde is an organic compound classified as carcinogenic since 2004 for nasopharyngeal cancer by the International Agency for Research on Cancer. Formol or Formaldehyde is, according to studies, very dangerous for health.

Why do we use 10% formalin?

Information about 10% Formalin – The fixative 10% buffered formalin is commonly used to preserve tissues for routine histology in many labs. The formaldehyde has a greater chance for oxidation in this concentration of tissue fixative and eventually the solution will start to drop in pH, in spite of the buffer.

We recommend that 10% buffered formalin solutions be used no longer than 3 months after they were initially mixed. The solution should be clear, colorless, with no precipitate and the pH should not be below 6.5. The other problem with 10% buffered formalin is the slowly increasing concentration of methanol (an unwanted byproduct of aging formaldehyde),

Methanol promotes clumping of proteins, instead of the cross-linking of proteins that formaldehyde performs. A methanol-free fixative will give the best preservation, particularly if you plan to use the tissue for antibody staining at a later time. The most common way to avoid methanol in a formaldehyde solution is to make the solution up fresh from crystalline paraformaldehyde.

  • Paraformaldehyde can be quite hazardous to handle and it is often difficult to get it to go into solution.
  • If your lab is not a regular user of formaldehyde fixatives, there are a couple of easier options that we recommend.
  • One option is to purchase methanol-free formaldehyde (aq) in sealed ampoules.
  • Simply add PBS to achieve the correct formaldehyde concentration and use immediately.

Ten 10 ml ampoules of 16% methanol-free formaldehyde costs approximately $27. The other option is to buy 10% neutral buffered Formalin (4% formaldehyde) from a scientific supply house, use it for 3-6 months and then discard it (as a hazardous material),

  • There will be some methanol in this solution (typically 1-2%), but if used soon after purchase this should not be significant for most users.
  • The buffered solution helps slow the acidification process.
  • A one liter bottle costs approximately $20-25.
  • Store the fixative at room temperature.
  • Please note, different suppliers use different buffering solutions.

For consistent immunohistochemistry and/or immunofluorescence, users should stick with one supplier.) Some labs have asked about using unbuffered 10% formalin. Unless there is a specific reason for this choice, we do not recommend it since this fixative rapidly becomes acidic.

What are the two main uses of formalin?

Uses of formaldehyde – When formaldehyde is dissolved in water, it is called formalin. This formalin is used as a disinfectant in industries, a preservative in some food products, funeral homes etc.

Used in industry Used in medicine Used in building and construction Food and other uses

What are the disadvantages of formalin?

Formaldehyde and Cancer Risk Formaldehyde is a colorless, flammable, strong-smelling chemical that is used in building materials and to produce many household products. It is used in pressed-wood products, such as particleboard, plywood, and fiberboard; glues and adhesives; permanent-press fabrics; paper product coatings; and certain insulation materials.

  1. In addition, formaldehyde is commonly used as an industrial,, and, and as a preservative in mortuaries and medical laboratories.
  2. Formaldehyde also occurs naturally in the environment.
  3. It is produced in small amounts by most living organisms as part of normal processes.
  4. According to a 1997 report by the U.S.

Consumer Product Safety Commission, formaldehyde is normally present in both indoor and outdoor air at low levels, usually less than 0.03 parts of formaldehyde per million parts of air (ppm). Materials containing formaldehyde can release formaldehyde gas or vapor into the air.

  1. One source of formaldehyde exposure in the air is automobile tailpipe emissions.
  2. During the 1970s, urea-formaldehyde foam insulation (UFFI) was used in many homes.
  3. However, few homes are now insulated with UFFI.
  4. Homes in which UFFI was installed many years ago are not likely to have high formaldehyde levels now.

Pressed-wood products containing formaldehyde resins are often a significant source of formaldehyde in homes. Other potential indoor sources of formaldehyde include cigarette smoke and the use of unvented fuel-burning appliances, such as gas stoves, wood-burning stoves, and kerosene heaters.

Industrial workers who produce formaldehyde or formaldehyde-containing products, laboratory technicians, certain health care professionals, and mortuary employees may be exposed to higher levels of formaldehyde than the general public. Exposure occurs primarily by formaldehyde gas or vapor from the air or by absorbing liquids containing formaldehyde through the skin.

When formaldehyde is present in the air at levels exceeding 0.1 ppm, some individuals may experience adverse effects such as watery eyes; burning sensations in the eyes, nose, and ; coughing; wheezing; ; and skin irritation. Some people are very sensitive to formaldehyde, whereas others have no reaction to the same level of exposure.

Although the short-term health effects of formaldehyde exposure are well known, less is known about its potential long-term health effects. In 1980, showed that exposure to formaldehyde could cause nasal cancer in rats. This finding raised the question of whether formaldehyde exposure could also cause cancer in humans.

In 1987, the U.S. Environmental Protection Agency (EPA) classified formaldehyde as a probable human carcinogen under conditions of unusually high or prolonged exposure (). Since that time, some studies of humans have suggested that formaldehyde exposure is associated with certain types of cancer.

The International Agency for Research on Cancer (IARC) classifies formaldehyde as a human carcinogen (). In 2011, the National Toxicology Program, an interagency program of the Department of Health and Human Services, named formaldehyde as a known human carcinogen in its 12 th Report on Carcinogens ().

Since the 1980s, the National Cancer Institute (NCI), a component of the National Institutes of Health (NIH), has conducted studies to determine whether there is an association between occupational exposure to formaldehyde and an increase in the risk of cancer.

  1. The results of this research have provided EPA and the Occupational Safety and Health Administration (OSHA) with information to evaluate the potential health effects of workplace exposure to formaldehyde.
  2. The long-term effects of formaldehyde exposure have been evaluated in epidemiologic studies (studies that attempt to uncover the patterns and causes of disease in groups of people).

One type of epidemiologic study is called a, A cohort is a group of people who may vary in their exposure to a particular factor, such as formaldehyde, and are followed over time to see whether they develop a disease. Another kind of epidemiologic study is called a,

Case-control studies begin with people who are diagnosed as having a disease (cases) and compare them to people without the disease (controls), trying to identify differences in factors, such as exposure to formaldehyde, that might explain why the cases developed the disease but the controls did not.

Several NCI surveys of professionals who are potentially exposed to formaldehyde in their work, such as anatomists and embalmers, have suggested that these individuals are at an increased risk of leukemia and brain cancer compared with the general population.

However, specific work practices and exposures were not characterized in these studies. An NCI case-control study among funeral industry workers that characterized exposure to formaldehyde also found an association between increasing formaldehyde exposure and from leukemia (). For this study, carried out among funeral industry workers who had died between 1960 and 1986, researchers compared those who had died from and lymphatic cancers and with those who died from other causes.

(Hematopoietic or such as leukemia develop in the blood or, Lymphatic cancers develop in the and that produce, store, and carry that fight and other diseases.) This analysis showed that those who had performed the most embalming and those with the highest estimated formaldehyde exposure had the greatest risk of myeloid leukemia.

There was no association with other cancers of the hematopoietic and lymphatic systems or with brain cancer. A number of cohort studies involving workers exposed to formaldehyde have recently been completed. One study, conducted by NCI, looked at 25,619 workers in industries with the potential for occupational formaldehyde exposure and estimated each worker’s exposure to the chemical while at work ().

The results showed an increased risk of death due to leukemia, particularly myeloid leukemia, among workers exposed to formaldehyde. This risk was associated with increasing peak and average levels of exposure, as well as with the duration of exposure, but it was not associated with,

An additional 10 years of data on the same workers were used in a study published in 2009 (). This analysis continued to show a possible link between formaldehyde exposure and cancers of the hematopoietic and lymphatic systems, particularly myeloid leukemia. As in the initial study, the risk was highest earlier in the follow-up period.

Risks declined steadily over time, such that the cumulative excess risk of myeloid leukemia was no longer at the end of the follow-up period. The researchers noted that similar patterns of risks over time had been seen for other agents known to cause leukemia.

  1. A cohort study of 11,039 textile workers performed by the National Institute for Occupational Safety and Health (NIOSH) also found an association between the duration of exposure to formaldehyde and leukemia deaths ().
  2. However, the evidence remains mixed because a cohort study of 14,014 British industry workers found no association between formaldehyde exposure and leukemia deaths ().

Formaldehyde undergoes rapid chemical changes immediately after absorption. Therefore, some scientists think that formaldehyde is unlikely to have effects at sites other than the upper, However, some laboratory studies suggest that formaldehyde may affect the lymphatic and hematopoietic systems.

  1. Based on both the epidemiologic data from cohort and case-control studies and the experimental data from laboratory research, NCI investigators have concluded that exposure to formaldehyde may cause leukemia, particularly myeloid leukemia, in humans.
  2. In addition, several case-control studies, as well as analysis of the large NCI industrial cohort (), have found an association between formaldehyde exposure and nasopharyngeal cancer, although some other studies have not.

Data from extended follow-up of the NCI cohort found that the excess of nasopharyngeal cancer observed in the earlier report persisted (). Earlier analysis of the NCI cohort found increased deaths among industrial workers compared with the general U.S.

  • Population.
  • However, the rate of lung cancer deaths did not increase with higher levels of formaldehyde exposure.
  • This observation led the researchers to conclude that factors other than formaldehyde exposure might have caused the increased deaths.
  • The most recent data on lung cancer from the cohort study did not find any relationship between formaldehyde exposure and lung cancer mortality.

In 1987, OSHA established a Federal standard that reduced the amount of formaldehyde to which workers can be exposed over an 8-hour workday from 3 ppm to 1 ppm. In May 1992, the standard was amended, and the formaldehyde exposure limit was further reduced to 0.75 ppm.

  • The EPA recommends the use of “exterior-grade” pressed-wood products to limit formaldehyde exposure in the home.
  • These products emit less formaldehyde because they contain resins, not urea resins.
  • Pressed-wood products include plywood, paneling, particleboard, and fiberboard and are not the same as pressure-treated wood products, which contain chemical preservatives and are intended for outdoor use.) Before purchasing pressed-wood products, including building materials, cabinetry, and furniture, buyers should ask about the formaldehyde content of these products.

Formaldehyde levels in homes can also be reduced by ensuring adequate ventilation, moderate temperatures, and reduced humidity levels through the use of air conditioners and dehumidifiers. The following organizations can provide additional resources that readers may find helpful: The EPA offers information about the use of formaldehyde in building materials and household products.

  • The EPA can be contacted at: U.S.
  • Environmental Protection Agency Office of Radiation and Indoor Air Indoor Environments Division Mail Code 6609J 1200 Pennsylvania Avenue, NW.
  • Washington, DC 20460 202–554–1404 (EPA Toxic Substance Control Act (TCSA) Assistance Line) The U.S.
  • Consumer Product Safety Commission (CPSC) has information about household products that contain formaldehyde.

CPSC can be contacted at: U.S. Consumer Product Safety Commission 4330 East West Highway Bethesda, MD 20814 1–800–638–2772 (1–800–638–CPSC) 301–595–7054 (TTY) The (FDA) maintains information about cosmetics and drugs that contain formaldehyde. FDA can be contacted at: U.S.

  • The Federal Emergency Management Agency (FEMA) has information about formaldehyde exposure levels in mobile homes and trailers supplied by FEMA after Hurricane Katrina. FEMA can be contacted at:
  • Federal Emergency Management Agency Post Office Box 10055 Hyattsville, MD 20782–7055 1–800–621–3362 (1–800–621–FEMA)
  • The Occupational Safety and Health Administration (OSHA) has information about occupational exposure limits for formaldehyde. OSHA can be contacted at:

U.S. Department of Labor Occupational Safety and Health Administration 200 Constitution Avenue Washington, DC 20210 1–800–321–6742 (1–800–321–OSHA) The National Toxicology Program (NTP) is an interagency program of the Department of Health and Human Services that was created to coordinate toxicology testing programs within the federal government; to develop and validate improved testing methods; and to provide information about potentially toxic chemicals to health, regulatory, and research agencies, scientific and medical communities, and the public.

  1. U.S. Environmental Protection Agency, Office of Air and Radiation. Report to Congress on Indoor Air Quality, Volume II: Assessment and Control of Indoor Air Pollution, 1989.
  2. International Agency for Research on Cancer (June 2004). IARC Monographs on the Evaluation of Carcinogenic Risks to Humans Volume 88 (2006): Formaldehyde, 2-Butoxyethanol and 1-tert-Butoxypropan-2-ol, Retrieved June 10, 2011, from:,
  3. National Toxicology Program (June 2011). Report on Carcinogens, Twelfth Edition. Department of Health and Human Services, Public Health Service, National Toxicology Program. Retrieved June 10, 2011, from:,
  4. Hauptmann M, Stewart PA, Lubin JH, et al. Mortality from lymphohematopoietic malignancies and brain cancer among embalmers exposed to formaldehyde. Journal of the National Cancer Institute 2009; 101(24):1696–1708.
  5. Hauptmann M, Lubin JH, Stewart PA, Hayes RB, Blair A. Mortality from lymphohematopoietic malignancies among workers in formaldehyde industries. Journal of the National Cancer Institute 2003; 95(21):1615–1623.
  6. Beane Freeman L, Blair A, Lubin JH, et al. Mortality from lymphohematopoietic malignancies among workers in formaldehyde industries: The National Cancer Institute Cohort. Journal of the National Cancer Institute 2009; 101(10):751–761.
  7. Pinkerton LE, Hein MJ, Stayner LT. Mortality among a cohort of garment workers exposed to formaldehyde: An update. Occupational Environmental Medicine 2004; 61:193–200.
  8. Coggon D, Harris EC, Poole J, Palmer KT. Extended follow-up of a cohort of British chemical workers exposed to formaldehyde. Journal of the National Cancer Institute 2003; 95(21):1608–1615.
  9. Hauptmann M, Lubin JH, Stewart PA, Hayes RB, Blair A. Mortality from solid cancers among workers in formaldehyde industries. American Journal of Epidemiology 2004; 159(12):1117–1130.

If you would like to reproduce some or all of this content, see for guidance about copyright and permissions. In the case of permitted digital reproduction, please credit the National Cancer Institute as the source and link to the original NCI product using the original product’s title; e.g., “Formaldehyde and Cancer Risk was originally published by the National Cancer Institute.” : Formaldehyde and Cancer Risk

What is the dilution ratio for formalin?

10% formalin is a 1:10 dilution of 100% formalin in water, i.e.1 part saturated formalde- hyde in water diluted with 9 parts plain water. Since 100% formalin contains 40% formaldehyde, a 1:10 dilution would contain 4% formaldehyde.

Can I use alcohol instead of formalin?

1. Introduction – Fixation is a physico-chemical phenomenon and reactions involving gradual diffusion of fixative into the tissues. It is an essential step in the evaluation and study of biopsy tissue specimens. It aids in the preservation of the tissue’s cellular architecture and composition during processing.

  • Fixation also maintains the spatial connection of proteins, carbohydrates, and other bioactive moieties to the cell, allowing them to be examined ( Woodyard, 2011 ).
  • For more than a century, the fixative of choice in routine histopathology has been a 10% solution of formalin (4% formaldehyde) diluted in water or in a buffered solution.
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Formalin maintains chemical activity and cellular antigenicity in tissues by forming covalent bonds between biological macromolecules ( Warmington et al., 2000 ). As a result, formalin-fixed paraffin-embedded (FFPE) tissues are now the most widely used fixatives in the world.

  1. However, formalin is not always the ‘gold standard’ fixative when other downstream biochemical and molecular analyses are considered ( Panzacci et al., 2019 ).
  2. In reality, the benefits of formalin as a histological preservation are outweighed by a variety of drawbacks, the most prominent of which are diminished immunohistochemical reactivity and rapid nucleic acid breakdown ( Cox et al., 2006, Moelans et al., 2011 ).

Although the degree of epitope modification differs with molecular targets, the crosslinking mechanism of formalin changes protein folding, decreasing the total availability of epitopes that can be bound by antibody ( Bogen et al., 2009, Hayat, 2000, O’Leary et al., 2009, Otali et al., 2009, Paavilainen et al., 2010 ).

Several proteolytic or heat-induced antigen retrieval methods have recently become commercially available to address this problem, restore normal protein folding, and improve the accessibility of epitope on fixed tissues ( Fowler et al., 2011, Paavilainen et al., 2010 ). Even though their mechanisms of action are unknown, these antigen retrieval techniques are currently considered the standard procedures for achieving high-quality staining.

As a result, antigen retrieval methods are often analytical and require high-level optimization. The baleful effects of formaldehyde have been scrutinized in public with all emotions and sensibility. Anatomists, technicians in histology and embalming laboratories, as well as medical students during their practical courses when they need to do the dissection of specimens, are all rigorously exposed to formaldehyde, which in certain cases reach to the threshold for inflammation of the eyes and upper respiratory tract.

There is no debate about the acute toxicity of formaldehyde and the incidence of contact dermatitis ( Pabst, 1987 ). The use of ethanol and methanol as a fixative, on the other hand, is a relatively non-toxic alternative to formalin. Ethanol and methanol are coagulating fixatives that break the hydrogen bonds to precipitate proteins.

While both ethanol and methanol have demonstrated their potential tissue fixation quality through routine use in cytological preparation, acting alone, both ethanol and methanol can cause tissue shrinkage and brittleness. Nucleic acid preservation as well as histological and immunohistochemical studies are possible with an alcohol-based fixative (molecular fixative, MF) ( Hostein et al., 2011 ).

  • Denaturation of the target antigens’ binding site can result in the use of an alcohol-based fixative.
  • As a result, antigen retrieval testing for each antibody is recommended, as it has been shown to produce a better immune reaction ( Magaki et al., 2019 ).
  • Milcheva et al.
  • 2013) found that after one hour of fixation in modified methacarn fixative, RNA extracted from the specimens still had intact rRNA subunits, while 24 h of exposure to alcohol-based fixative resulted in total loss of rRNA subunit detection on an agarose gel.

Alcohol-based tissue fixatives, such as ethanol and methanol, are more efficient preserving agents than buffered formalin. These fixatives allowed the recovery of higher quality DNA by controlling dehydration rather than cross-linking ( Duval et al., 2010 ).

Alcohol-based fixatives work by precipitating proteins, which does not disguise their antigenicity and eliminates the need for antigen retrieval on slides. It has long been recognized that alcohol supplemented with acetic acid results in strong tissue morphology preservation ( Buchwalow et al., 2010 ).

Both methanol and ethanol combined with glacial acetic acid, with either routine or manual processing of the tissue specimen, provide an excellent and reliable tool for research studies that aim to investigate tissue morphology, immunohistochemical detection of proteins, and molecular analyses in normal and pathologically changed tissues, particularly when tissue morphology, immunohistochemical detection of proteins, and molecular analyses in normal and pathologically changed tissues are investigated.

What happens if you accidentally touch formalin?

1910.1048 App A – Substance Technical Guidelines for Formalin | Occupational Safety and Health Administration Appendix A to § 1910.1048 – Substance Technical Guidelines for Formalin The following Substance Technical Guideline for Formalin provides information on uninhibited formalin solution (37% formaldehyde, no methanol stabilizer).

  1. It is designed to inform employees at the production level of their rights and duties under the formaldehyde standard whether their job title defines them as workers or supervisors.
  2. Much of the information provided is general; however, some information is specific for formalin.
  3. When employee exposure to formaldehyde is from resins capable of releasing formaldehyde, the resin itself and other impurities or decomposition products may also be toxic, and employers should include this information as well when informing employees of the hazards associated with the materials they handle.

The precise hazards associated with exposure to formaldehyde depend both on the form (solid, liquid, or gas) of the material and the concentration of formaldehyde present. For example, 37-50 percent solutions of formaldehyde present a much greater hazard to the skin and eyes from spills or splashes than solutions containing less than 1 percent formaldehyde.

  • Substance Identification
  • Chemical Name: Formaldehyde
  • Chemical Family: Aldehyde
  • Chemical Formula: HCHO
  • Molecular Weight: 30.03
  • Chemical Abstracts Service Number (CAS Number): 50-00-0
  • Synonyms: Formalin; Formic Aldehyde; Paraform; Formol; Formalin (Methanol-free); Fyde; Formalith; Methanal; Methyl Aldehyde; Methylene Glycol; Methylene Oxide; Tetraoxymethalene; Oxomethane; Oxymethylene
  • Components and Contaminants
  • Percent: 37.0 Formaldehyde
  • Percent: 63.0 Water
  • (Note – Inhibited solutions contain methanol.)
  • Other Contaminants: Formic acid (alcohol free)
  • Exposure Limits:
  • OSHA TWA – 0.75 ppm
  • OSHA STEL – 2 ppm
  • Physical Data
  • Description: Colorless liquid, pungent odor
  • Boiling point: 214 °F (101 °C)
  • Specific Gravity: 1.08 (H 2 O = 1 @ 20 °C)

pH: 2.8-4.0

  1. Solubility in Water: Miscible
  2. Solvent Solubility: Soluble in alcohol and acetone
  3. Vapor Density: 1.04 (Air = 1 @ 20 °C)
  4. Odor Threshold: 0.8-1 ppm
  5. Fire and Explosion Hazard
  6. Moderate fire and explosion hazard when exposed to heat or flame.
  7. The flash point of 37% formaldehyde solutions is above normal room temperature, but the explosion range is very wide, from 7 to 73% by volume in air.
  8. Reaction of formaldehyde with nitrogen dioxide, nitromethane, perchloric acid and aniline, or peroxyformic acid yields explosive compounds.
  9. Flash Point: 185 °F (85 °C) closed cup
  10. Lower Explosion Limit: 7%
  11. Upper Explosion Limit: 73%
  12. Autoignition Temperature: 806 °F (430 °C)
  13. Flammability (OSHA): Category 4 flammable liquid

Extinguishing Media: Use dry chemical, “alcohol foam”, carbon dioxide, or water in flooding amounts as fog. Solid streams may not be effective. Cool fire-exposed containers with water from side until well after fire is out. Use of water spray to flush spills can also dilute the spill to produce nonflammable mixtures.

  1. Water runoff, however, should be contained for treatment.
  2. National Fire Protection Association Section 325M Designation: Health: 2 – Materials hazardous to health, but areas may be entered with full-faced mask self-contained breathing apparatus which provides eye protection.
  3. Flammability: 2 – Materials which must be moderately heated before ignition will occur.

Water spray may be used to extinguish the fire because the material can be cooled below its flash point. Reactivity: D – Materials which (in themselves) are normally stable even under fire exposure conditions and which are not reactive with water. Normal fire fighting procedures may be used.

  1. Reactivity Stability: Formaldehyde solutions may self-polymerize to form paraformaldehyde which precipitates.
  2. Incompatibility (Materials to Avoid): Strong oxidizing agents, caustics, strong alkalies, isocyanates, anhydrides, oxides, and inorganic acids.
  3. Formaldehyde reacts with hydrochloric acid to form the potent carcinogen, bis-chloromethyl ether.

Formaldehyde reacts with nitrogen dioxide, nitromethane, perchloric acid and aniline, or peroxyformic acid to yield explosive compounds. A violent reaction occurs when formaldehyde is mixed with strong oxidizers. Hazardous Combustion or Decomposition Products: Oxygen from the air can oxidize formaldehyde to formic acid, especially when heated.

Formic acid is corrosive. Health Hazard Data Acute Effects of Exposure Ingestion (Swallowing): Liquids containing 10 to 40% formaldehyde cause severe irritation and inflammation of the mouth, throat, and stomach. Severe stomach pains will follow ingestion with possible loss of consciousness and death.

Ingestion of dilute formaldehyde solutions (0.03-0.04%) may cause discomfort in the stomach and pharynx. Inhalation (Breathing): Formaldehyde is highly irritating to the upper respiratory tract and eyes. Concentrations of 0.5 to 2.0 ppm may irritate the eyes, nose, and throat of some individuals.

Concentrations of 3 to 5 ppm also cause tearing of the eyes and are intolerable to some persons. Concentrations of 10 to 20 ppm cause difficulty in breathing, burning of the nose and throat, cough, and heavy tearing of the eyes, and 25 to 30 ppm causes severe respiratory tract injury leading to pulmonary edema and pneumonitis.

A concentration of 100 ppm is immediately dangerous to life and health. Deaths from accidental exposure to high concentrations of formaldehyde have been reported. Skin (Dermal): Formalin is a severe skin irritant and a sensitizer. Contact with formalin causes white discoloration, smarting, drying, cracking, and scaling.

Prolonged and repeated contact can cause numbness and a hardening or tanning of the skin. Previously exposed persons may react to future exposure with an allergic eczematous dermatitis or hives. Eye Contact: Formaldehyde solutions splashed in the eye can cause injuries ranging from transient discomfort to severe, permanent corneal clouding and loss of vision.

The severity of the effect depends on the concentration of formaldehyde in the solution and whether or not the eyes are flushed with water immediately after the accident. Note. The perception of formaldehyde by odor and eye irritation becomes less sensitive with time as one adapts to formaldehyde.

  • Acute Animal Toxicity:
  • Oral, rats: LD50 = 800 mg/kg
  • Oral, mouse: LD50 = 42 mg/kg
  • Inhalation, rats: LCLo = 250 mg/kg
  • Inhalation, mouse: LCLo = 900 mg/kg
  • Inhalation, rats: LC50 = 590 mg/kg
  • Chronic Effects of Exposure

Carcinogenicity: Formaldehyde has the potential to cause cancer in humans. Repeated and prolonged exposure increases the risk. Various animal experiments have conclusively shown formaldehyde to be a carcinogen in rats. In humans, formaldehyde exposure has been associated with cancers of the lung, nasopharynx and oropharynx, and nasal passages.

Mutagenicity: Formaldehyde is genotoxic in several in vitro test systems showing properties of both an initiator and a promoter. Toxicity: Prolonged or repeated exposure to formaldehyde may result in respiratory impairment. Rats exposed to formaldehyde at 2 ppm developed benign nasal tumors and changes of the cell structure in the nose as well as inflamed mucous membranes of the nose.

Structural changes in the epithelial cells in the human nose have also been observed. Some persons have developed asthma or bronchitis following exposure to formaldehyde, most often as the result of an accidental spill involving a single exposure to a high concentration of formaldehyde.

Emergency and First Aid Procedures Ingestion (Swallowing): If the victim is conscious, dilute, inactivate, or absorb the ingested formaldehyde by giving milk, activated charcoal, or water. Any organic material will inactivate formaldehyde. Keep affected person warm and at rest. Get medical attention immediately.

If vomiting occurs, keep head lower than hips. Inhalation (Breathing): Remove the victim from the exposure area to fresh air immediately. Where the formaldehyde concentration may be very high, each rescuer must put on a self-contained breathing apparatus before attempting to remove the victim, and medical personnel should be informed of the formaldehyde exposure immediately.

  1. If breathing has stopped, give artificial respiration.
  2. Eep the affected person warm and at rest.
  3. Qualified first-aid or medical personnel should administer oxygen, if available, and maintain the patient’s airways and blood pressure until the victim can be transported to a medical facility.
  4. If exposure results in a highly irritated upper respiratory tract and coughing continues for more than 10 minutes, the worker should be hospitalized for observation and treatment.

Skin Contact: Remove contaminated clothing (including shoes) immediately. Wash the affected area of your body with soap or mild detergent and large amounts of water until no evidence of the chemical remains (at least 15 to 20 minutes). If there are chemical burns, get first aid to cover the area with sterile, dry dressing, and bandages.

Get medical attention if you experience appreciable eye or respiratory irritation. Eye Contact: Wash the eyes immediately with large amounts of water occasionally lifting lower and upper lids, until no evidence of chemical remains (at least 15 to 20 minutes). In case of burns, apply sterile bandages loosely without medication.

Get medical attention immediately. If you have experienced appreciable eye irritation from a splash or excessive exposure, you should be referred promptly to an opthamologist for evaluation. Emergency Procedures Emergencies: If you work in an area where a large amount of formaldehyde could be released in an accident or from equipment failure, your employer must develop procedures to be followed in event of an emergency.

  • You should be trained in your specific duties in the event of an emergency, and it is important that you clearly understand these duties.
  • Emergency equipment must be accessible and you should be trained to use any equipment that you might need.
  • Formaldehyde contaminated equipment must be cleaned before reuse.

If a spill of appreciable quantity occurs, leave the area quickly unless you have specific emergency duties. Do not touch spilled material. Designated persons may stop the leak and shut off ignition sources if these procedures can be done without risk.

Designated persons should isolate the hazard area and deny entry except for necessary people protected by suitable protective clothing and respirators adequate for the exposure. Use water spray to reduce vapors. Do not smoke, and prohibit all flames or flares in the hazard area. Special Firefighting Procedures: Learn procedures and responsibilities in the event of a fire in your workplace.

Become familiar with the appropriate equipment and supplies and their location. In firefighting, withdraw immediately in case of rising sound from venting safety device or any discoloration of storage tank due to fire. Spill, Leak, and Disposal Procedures Occupational Spill: For small containers, place the leaking container in a well ventilated area.

Take up small spills with absorbent material and place the waste into properly labeled containers for later disposal. For larger spills, dike the spill to minimize contamination and facilitate salvage or disposal. You may be able to neutralize the spill with sodium hydroxide or sodium sulfite. Your employer must comply with EPA rules regarding the clean-up of toxic waste and notify state and local authorities, if required.

If the spill is greater than 1,000 lb/day, it is reportable under EPA’s Superfund legislation. Waste Disposal: Your employer must dispose of waste containing formaldehyde in accordance with applicable local, state, and Federal law and in a manner that minimizes exposure of employees at the site and of the clean-up crew.

  1. Monitoring and Measurement Procedures Monitoring Requirements: If your exposure to formaldehyde exceeds the 0.5 ppm action level or the 2 ppm STEL, your employer must monitor your exposure.
  2. Your employer need not measure every exposure if a “high exposure” employee can be identified.
  3. This person usually spends the greatest amount of time nearest the process equipment.

If you are a “representative employee”, you will be asked to wear a sampling device to collect formaldehyde. This device may be a passive badge, a sorbent tube attached to a pump, or an impinger containing liquid. You should perform your work as usual, but inform the person who is conducting the monitoring of any difficulties you are having wearing the device.

Evaluation of 8-hour Exposure: Measurements taken for the purpose of determining time-weighted average (TWA) exposures are best taken with samples covering the full shift. Samples collected must be taken from the employee’s breathing zone air. Short-term Exposure Evaluation: If there are tasks that involve brief but intense exposure to formaldehyde, employee exposure must be measured to assure compliance with the STEL.

Sample collections are for brief periods, only 15 minutes, but several samples may be needed to identify the peak exposure. Monitoring Techniques: OSHA’s only requirement for selecting a method for sampling and analysis is that the methods used accurately evaluate the concentration of formaldehyde in employees’ breathing zones.

Sampling and analysis may be performed by collection of formaldehyde on liquid or solid sorbents with subsequent chemical analysis. Sampling and analysis may also be performed by passive diffusion monitors and short-term exposure may be measured by instruments such as real-time continuous monitoring systems and portable direct reading instruments.

Notification of Results: Your employer must inform you of the results of exposure monitoring representative of your job. You may be informed in writing, but posting the results where you have ready access to them constitutes compliance with the standard.

  1. Protective Equipment and Clothing Respiratory Protection: Use NIOSH-approved full facepiece negative pressure respirators equipped with approved cartridges or canisters within the use limitations of these devices.
  2. Present restrictions on cartridges and canisters do not permit them to be used for a full workshift.) In all other situations, use positive pressure respirators such as the positive-pressure air purifying respirator or the self-contained breathing apparatus (SCBA).

If you use a negative pressure respirator, your employer must provide you with fit testing of the respirator at least once a year. Protective Gloves: Wear protective (impervious) gloves provided by your employer, at no cost, to prevent contact with formalin.

  • Your employer should select these gloves based on the results of permeation testing and in accordance with the ACGIH Guidelines for Selection of Chemical Protective Clothing.
  • Eye Protection: If you might be splashed in the eyes with formalin, it is essential that you wear goggles or some other type of complete protection for the eye.

You may also need a face shield if your face is likely to be splashed with formalin, but you must not substitute face shields for eye protection. (This section pertains to formaldehyde solutions of 1% or more.) Other Protective Equipment: You must wear protective (impervious) clothing and equipment provided by your employer at no cost to prevent repeated or prolonged contact with formaldehyde liquids.

  • If you are required to change into whole-body chemical protective clothing, your employer must provide a change room for your privacy and for storage of your normal clothing.
  • If you are splashed with formaldehyde, use the emergency showers and eyewash fountains provided by your employer immediately to prevent serious injury.

Report the incident to your supervisor and obtain necessary medical support. Entry Into an IDLH Atmosphere Enter areas where the formaldehyde concentration might be 100 ppm or more only with complete body protection including a self-contained breathing apparatus with a full facepiece operated in a positive pressure mode or a supplied air respirator with full facepiece and operated in a positive pressure mode.

This equipment is essential to protect your life and health under such extreme conditions. Engineering Controls Ventilation is the most widely applied engineering control method for reducing the concentration of airborne substances in the breathing zones of workers. There are two distinct types of ventilation.

Local Exhaust: Local exhaust ventilation is designed to capture airborne contaminants as near to the point of generation as possible. To protect you, the direction of contaminant flow must always be toward the local exhaust system inlet and away from you.

  • General (Mechanical): General dilution ventilation involves continuous introduction of fresh air into the workroom to mix with the contaminated air and lower your breathing zone concentration of formaldehyde.
  • Effectiveness depends on the number of air changes per hour.
  • Where devices emitting formaldehyde are spread out over a large area, general dilution ventilation may be the only practical method of control.

Work Practices: Work practices and administrative procedures are an important part of a control system. If you are asked to perform a task in a certain manner to limit your exposure to formaldehyde, it is extremely important that you follow these procedures.

Medical Surveillance Medical surveillance helps to protect employees’ health. You are encouraged strongly to participate in the medical surveillance program. Your employer must make a medical surveillance program available at no expense to you and at a reasonable time and place if you are exposed to formaldehyde at concentrations above 0.5 ppm as an 8-hour average or 2 ppm over any 15-minute period.

You will be offered medical surveillance at the time of your initial assignment and once a year afterward as long as your exposure is at least 0.5 ppm (TWA) or 2 ppm (STEL). Even if your exposure is below these levels, you should inform your employer if you have signs and symptoms that you suspect, through your training, are related to your formaldehyde exposure because you may need medical surveillance to determine if your health is being impaired by your exposure.

  1. The surveillance plan includes:
  2. (a) A medical disease questionnaire.
  3. (b) A physical examination if the physician determines this is necessary.
  4. If you are required to wear a respirator, your employer must offer you a physical examination and a pulmonary function test every year.

The physician must collect all information needed to determine if you are at increased risk from your exposure to formaldehyde. At the physician’s discretion, the medical examination may include other tests, such as a chest x-ray, to make this determination.

After a medical examination the physician will provide your employer with a written opinion which includes any special protective measures recommended and any restrictions on your exposure. The physician must inform you of any medical conditions you have which would be aggravated by exposure to formaldehyde.

All records from your medical examinations, including disease surveys, must be retained at your employer’s expense. Emergencies If you are exposed to formaldehyde in an emergency and develop signs or symptoms associated with acute toxicity from formaldehyde exposure, your employer must provide you with a medical examination as soon as possible.

What is the difference between formaldehyde and formalin?

Formaldehyde is a small molecule that is a gas at normal conditions. Formalin is ~40% saturated solution of formaldehyde. In this solution, formaldehyde forms polymers of paraformaldehyde.10% formalin is a 1:10 dilution of formalin and contains ~4% paraformaldehyde (or formaldehyde).

Is there a difference between formaldehyde and formalin?

Formaldehyde solution CAS No.

What happens when formalin is heated?

The thermal decomposition of formaldehyde into carbon monoxide and hydrogen has been shown to be a homogeneous bimolecular reaction over the pressure range 30-400 mm. Simultaneous condensation reactions occur. The decomposition has been studied from 510°-607° C, and the energy of activation found to be 44,500 calories.

What is the best way to preserve fish through drying method?

Common Ways to Dry Fish – When using this method of fish preservation, you want to use the freshest fish possible to get the best results. The easiest way to get started is by using very salty brine, three parts water to one part salt. By brining the fish, you remove the blood from the meat.

Another traditional method of fish preservation involves dry-salting, which is exactly what it sounds like. After cleaning the fish, you lay them in a dry basket and cover them with salt. Use about one-third of the weight of the fish in salt. Cover the basket and let it sit for 9-10 days. The salt will draw all of the moisture out of the fish, and starve bacteria so they won’t be able to grow.

Modern technology means you can make fish this way in a dehydrator or even make smoked fish using a charcoal smoker, Read the instruction manual carefully before trying this. And make sure to wash all parts of the dehydrator in hot soapy water before and after preparing the fish.

How do you spray formalin?

Description – Formalin is an aldehyde disinfectant product used extensively in livestock housing. Formalin can be applied as a wet spray to surfaces and equipment however it is generally applied as a thermal fog (thermal fogging) when disinfecting large areas such as poultry houses.

  • Formalin is also used as a foot disinfectant for sheep and cattle.
  • If using as a spray dilute 1 part formalin to 4 parts water (1:4) and apply at 400 ml per square metre of surface area.
  • When thermal fogging formalin is used undiluted at a rate of 10 litres per 10,000 square feet (approximately 1,000 square metres) of floor space.

Apply using a suitable thermal fog machine. Formalin 38/10 (38% formaldehyde). CAS 50-00-0. EC Number 200-001-8. UN2209. Download the Formalin safety data sheet,