Blood Disposing and Handling

Michael LavoieVeterinary Assistant ProgramMiddlesex Community College12/19/12

Blood Vessels

1. Arteries

--carry blood away from the heart

--usually spurt blood when cut

--all except the pulmonary artery carry oxygenated blood

--thick walled and elastic

pulse: expansion and contraction of the artery walls in response to the heartbeat

Veins

--carry blood toward the heart --contain valves --closer to the body surface than the arteries --all except the pulmonary vein carry

deoxygenated blood --thinner, less muscular and elastic than

arteries --depend upon muscle and diaphragm

movements for blood flow

Capillaries

--most numerous vessels

--connect arteries to veins

--microscopic, one cell thick walls

--site of much exchange between the blood and the intracellular fluid (lymph) by diffusion

Blood Composition

Plasma 55% (liquid part of the blood); Blood Cells 45%

The Parts of Blood

1. Plasma =carries everything

2. Red Blood Cells =(RBC) gas exchange

3. White blood Cells =(WBC) fight infection

4. Platelets = clotting

55% plasma

45 %

RBC, WBC and platelets

Plasma- nonliving

Yellow liquid (92% H2O) 8 % nutrients, salts, urea, hormones Carries:

RBC, WBC, Platelets, Carbon dioxide, food and waste

BLOOD CELL TYPES

Red Blood Cells: most numerous biconcave disc shaped smaller than white blood cells, larger than platelets no nucleus when mature produced in the red marrow of long bones destroyed in the liver and spleen contain the iron protein compound HEMOGLOBIN

whose chief function is to combine with oxygen and carry it to the cells

Red Blood Cells- living 5 million in 1 drop of blood (most common) Shape = donut

Draw here:

Made in bone marrow Live approximately 120-125 days

Hemoglobin = oxygen containing pigment Binds to oxygen and carries it to the cells Gives red blood cells its red color

White blood cells- living AKA- Lymphocytes or Leukocytes White blood cells are larger than red blood cells,

but there are less of them. 8000 in one drop of blood

Function of White Blood Cells surround and digest bacteria Attack bacteria and viruses

**Pus = WBC + dead bacteria

YouTube - White Blood Cell Chases Bacteria

White Blood cells

--largest blood cells--several different types

--about 8,000 per drop of blood

--most are formed in the bone marrow or in the lymph tissue

--most protect the body against diseases by forming antibodies or engulfing bacteria

Main White Blood Cell Types

1. Phagocytes-- engulf bacteria and viruses by phagocytosis

--able to leave the bloodstream

and move between the cells of

the body by squeezing through

the capillary walls

2. Lymphocytes--produce antibodies which clump bacterial poisons or bacteria (antigens) (antigens--foreign substances in the body)

Platelets- living Bits of cells Live for approximately 10 days

Function of Platelets creates fibrin = enzyme that helps clot blood

(tiny threads seal cuts)

3. Platelets

--smallest blood cells (fragments)

--150,000 to 300,000 per drop of blood

--needed for clotting

Clotting:

Involves a series of enzyme controlled reactions resulting in the formation of protein fibers that trap blood cells and form a clot.

Restraint

Dogs, Cats, and Sheep: usually require only physical restraint to collect blood.

Swine: may require only physical restraint if they have been trained to the procedure. Otherwise, chemical restraint or sedation is recommended.

Rabbits, Mice and Rats: may be placed in appropriate restraining devices or chemically restrained with anesthetics or sedatives.

Birds: usually require only physical restraint to collect blood.

Blood Drawing

Cats and dogs can have blood drawn in many ways Jugular Cephalic Saphenous veins

Dogs- Cephalic

Blood samples can be collected from the cephalic vein which is superficial and easily accessible.

The technique is quick and appropriate for taking small blood samples.

Blood is collected from the forelimb, starting distally (near the paw) and working proximally (away from paw).

Where multiple samples are taken, the use of the forelegs should be alternated

Cephalic

Aseptic technique should be used, including clipping the hair around the sampling site.

Blood should not be taken from a vein that has been used to administer intravenous drugs.

Volume

3 - 5 ml of blood can be collected per sample and, depending on sample volume.

Normally no more than eight samples in a 24-hour period; more frequent sampling can cause the forelegs to become bruised.

The number of attempts to obtain blood should be minimized to a maximum of three needle sticks for each sample.

Bleeding should be stopped, before the dog is returned to its pen, by finger pressure to the sampling site for approximately 30 seconds.

Jugular Vein Restraint

The dog is usually in the sitting position. The restrainer elevates head with one hand

wrapped around muzzle and the nose directed upward. 

The front legs may also need to be controlled. 

The person taking blood in this case will hold off the jugular vein with one hand and draw the blood with the other hand.

The dog is restrained manually. Sampling sites are alternated between the

two jugular veins, starting distally at the base of the neck and moving towards the head along the jugular groove.

The vein is raised by compressing it just dorsal to the thoracic inlet, ventral to the venepuncture site.

Aseptic technique should be used, including clipping the hair around the sampling site.

2- 20 ml of blood can be taken per sample and, depending on sample volume, up to eight samples in a 24-hour period; more frequent sampling can cause bruising.

The number of attempts to obtain blood should be minimized to a maximum of three needle sticks for each sample.

Bleeding should be stopped, before the dog is returned to its pen, by finger pressure to the sampling site for approximately 30 seconds.

Saphenous Vein Draw

The vein is located on the lateral hind leg above the hock joint, this vein collapses easier than others  and is used as a last resort usually. 

The patient may be standing or lying on it's side for the venipuncture. 

The vein is held off at or slightly below the stifle joint  by wrapping one hand around the leg and applying pressure, the other hand and arm are used to restrain the patient.

The person taking the blood steadies the lower leg and does the venipuncture with their other hand.

3 - 5 ml of blood can be collected per sample and, depending on sample volume.

Normally no more than eight samples in a 24-hour period; more frequent sampling can cause the forelegs to become bruised.

The number of attempts to obtain blood should be minimized to a maximum of three needle sticks for each sample.

Bleeding should be stopped, before the dog is returned to its pen, by finger pressure to the sampling site for approximately 30 seconds.

Jugular on Cats

The jugular vein maybe the vein of choice depending on future medical procedures or the condition of other veins.

It is the largest peripheral vein.  The cat is usually positioned over the edge of the

table, a towel can be placed under the cat for comfort.

The restrainer is positioned behind and to the side of the patient. The front legs are held together (split finger position) and pulled downward with one hand.

The head is held and positioned with the other hand. The position of the head is very important so that

the jugular vein is adequately exposed. The hand is wrapped around the head to keep the

mouth closed and the nose is pointed upward.  The jugular venipuncture can also be performed

with the cat in dorsal recumbency (lying on it's stomach) in this case the cat is usually restrained in a bag or towel.

The cat is restrained manually. Sampling sites are alternated between the

two jugular veins, starting distally at the base of the neck and moving towards the head along the jugular groove.

The vein is raised by compressing it just dorsal to the thoracic inlet, ventral to the venepuncture site.

Aseptic technique should be used, including clipping the hair around the sampling site.

3- 12 ml of blood can be taken per sample and, depending on sample volume, up to eight samples in a 24-hour period; more frequent sampling can cause bruising.

The number of attempts to obtain blood should be minimized to a maximum of three needle sticks for each sample.

Bleeding should be stopped, before the cat is returned to its pen, by finger pressure to the sampling site for approximately 30 seconds.

Cephalic Vein in Cats

Feline cephalic venipuncture is very similar to that of the dog.

The cat is placed in ventral recumbency (it can be in a bag or towel if needed).

A front leg is extended and the vein rolled outward and pressure applied to hold it off.

The head is controlled with the other hand and back legs controlled by the restrainer's body.

Blood samples can be collected from the cephalic vein which is superficial and easily accessible.

The technique is quick and appropriate for taking small blood samples.

Blood is collected from the forelimb, starting distally (near the paw) and working proximally (away from paw).

Where multiple samples are taken, the use of the forelegs should be alternated

Aseptic technique should be used, including clipping the hair around the sampling site.

Blood should not be taken from a vein that has been used to administer intravenous drugs.

3 - 5 ml of blood can be collected per sample and, depending on sample volume.

Normally no more than eight samples in a 24-hour period; more frequent sampling can cause the forelegs to become bruised.

The number of attempts to obtain blood should be minimized to a maximum of three needle sticks for each sample.

Bleeding should be stopped, before the dog is returned to its pen, by finger pressure to the sampling site for approximately 30 seconds.

Femoral Blood draw on Cats

The femoral vein is located on the inside of the hind leg.

It is quite easily palpated and  superficial to the artery.

This vein is easily collapsed and easily double punctured. 

It is used for intravenous injection and small volume blood collection.

The cat is placed in lateral recumbency (usually in the stretch or scruff and stretch positions).

Blood is taken from the down leg (on the table). The "up leg" and tail are held out of the way. The heel of the hand holding the "up" leg is placed

in the groin area to occlude the femoral vein (pressure is applied at the femoral triangle proximal to venipuncture site).

The restrainer holds the scruff and holds off the vein, the venipuncture "down" leg is held by the person taking the blood.

3 - 5 ml of blood can be collected per sample and, depending on sample volume.

Normally no more than eight samples in a 24-hour period; more frequent sampling can cause the forelegs to become bruised.

The number of attempts to obtain blood should be minimized to a maximum of three needle sticks for each sample.

Bleeding should be stopped, before the dog is returned to its pen, by finger pressure to the sampling site for approximately 30 seconds.

Mouse and rat Blood sampling The lateral tail vein is usually used and 50 ul to 0.2 ml of blood

can be obtained per sample depending on the size of the animal and specific requirements.

.1-2ml of blood can be obtained from rats The tail may need to be washed with diluted Hibiscrub (1%) in

order to see the blood vessel. To avoid bruising and damage to the tail, normally no more than

two blood samples should be taken per session and in any one 24-hour period.

The number of attempts to take a blood sample should be minimized (no more than three needle sticks in any one attempt) and sufficient time should be given for the tail to recover between blood sampling sessions.

Alternate sides of the tail should be used and successive needle punctures moved towards the tail base.

Tail vein sampling is suitable for all strains but is more difficult in black or pigmented mice. 

t is quick and simple to perform for competent individuals. 

This technique may require the animals to be warmed in order to dilate the blood vessel prior to taking the sample.

This may be stressful and can cause dehydration due to salivation, in addition to increasing metabolic rate, which may affect the experimental data.

Saphenous vein draw

To collect blood, the hind leg should be immobilized in the extended position by applying gentle downward pressure immediately above the knee joint.

This stretches the skin over the ankle, making it easier to shave, and immobilizes the saphenous vein. 

Aseptic technique should be used.

Hamster Blood Draw: Saphenous Conscious hamsters should be restrained

either manually or using a restraint tube. This can cause stress and therefore the

duration of restraint should be minimized. Where a restraint tube is used, it should be

appropriate for the size of the hamster. All forms of restraining equipment should be

frequently washed to prevent pheromonally-induced stress or cross-infection.

Saphenous vein

To collect blood, the hind leg, should be immobilized in the extended position by applying gentle downward pressure immediately above the knee joint. 

This stretches the skin over the ankle, making it easier to shave, and immobilizes the saphenous vein.

Aseptic technique should be used.

No more than a maximum of four blood samples should be taken within any 24-hour period, depending on sample volume.

A single sample equivalent to 0.5% of the animal's bodyweight, which can usually be repeated at fortnightly intervals without disturbances to its haematological status.

Alternatively, daily samples corresponding to 0.05% bodyweight.

Adverse effects: Bruising Haemorrhage Infection Temporary favoring of the limb

Guinea Pig Blood draws: Saphenous Conscious guinea pigs should be restrained

either manually or using a restraint tube. This can cause stress and therefore the

duration of restraint should be minimized. Where a restraint tube is used, it should be

appropriate to the size of the animal. All forms of restraining equipment should be

frequently washed to prevent pheromonally-induced stress or cross-infection.

Blood is collected from the lateral saphenous vein, which runs dorsally and then laterally over the tarsal joint.

To collect blood, the hind leg, should be immobilised in the extended position by applying gentle downward pressure immediately above the knee joint. 

Aseptic technique should be used

No more than four blood samples should be taken within any 24-hour period, depending on sample volume.

A single sample equivalent to 0.5% of the animal's bodyweight, which can usually be repeated at fort nightly intervals without disturbances to its haematological status.

Alternatively, daily samples corresponding to 0.05% bodyweight.

Rabbit Blood draw: Marginal ear vein/artery Removal of blood from the marginal ear vein

or artery is one of the most common and least invasive methods of talking blood from a rabbit.

This technique can be used with all strains and for single and repeat samples.

Use of the artery is normally used for larger volume samples or for arterial blood, but carries a greater risk of haematoma and bruising.

The rabbit should be restrained and it can be helpful to wrap the animal in a large cloth to avoid inadvertent movement.

Restraint can cause stress, therefore the duration of restraint should be minimized.

Blood is taken from the tip of the ear, away from the base of the ear.

Serial blood samples can be taken by moving towards the base of the ear on the same vein and by alternating ears.

The ear should be warmed in order to dilate the vessel.

Depending on the size of the rabbit and the frequency of sampling, 0.5 - 10 ml of blood can be collected.

Up to eight samples can be collected in any 24-hour period, depending on sample volume

In order to minimize damage to the ear the number of attempts to take a blood sample should be minimized (no more than three needle sticks in any one attempt).

Drawing blood on pigs: Ear vein Pigs are intelligent and can be trained to accept

handling and restraint. They will remember receiving a reward (e.g. food

treat) after the procedure, which can make them easier to handle on subsequent occasions.

Removal of blood from the marginal ear veins of pigs is suitable for single and repeat sampling of small volumes (<1 ml).

The technique can be used with all breeds, although the ear veins of minipigs are small and can collapse if too much vacuum is applied when withdrawing the sample.

The pig needs to be restrained for sampling and this can be stressful.

Stress can be minimized by training the animal to co-operate with the procedure and by conducting it in a quite environment, if possible.

Sedation of the pig should be considered, particularly when withdrawing larger blood volumes.

Large pigs can be bled whilst standing and restrained by a snout rope.

Minipigs and small pigs can be held across the lap or against the body.

The vein is occluded at the base of the lateral surface of the ear.

The needle is slid towards the base of the ear. When the vein has been punctured, the emerging blood

can be collected directly by capillary action into appropriate tubes.

Serial blood samples can be taken by moving towards the base of the ear on the same vein and by alternating ears.

Blood flow should be stopped before the animal is retuned to its pen by applying finger pressure on soft tissue placed at the blood sampling site for approximately 2 minutes.

What is Medical Waste?

Medical Waste includes all infectious hazardous wastes, (including low-level radioactive wastes) and any other wastes that is generated from all types of health care facilities. These types of health care facilities include:

Hospitals Clinics Doctor’s offices (veterinary and human) Medical laboratories

What is regulated medical waste? Regulated medical waste is a subset of all medical

wastes and includes seven different categories:1. Cultures and stocks of infectious agents

2. Human pathological wastes (e.g. tissues, body parts)

3. Human and animal blood and blood products

4. Sharps (e.g. hypodermic needles and syringes)

5. Certain animal wastes

6. Certain isolation wastes (wastes from patients with highly communicable diseases)

7. Unused sharps

What is an infectious waste?

There are several factors that define infectious waste

1. Presence of pathogen of sufficient virulence2. Dose3. Portal of entry4. Resistance of the host It must contain a pathogen with sufficient

virulence and quantity so that the exposure to the waste by a person or animal could result in an infectious disease

What is an infectious waste?

1. Isolation wastes: wastes generated by hospital patients who are isolated to protect others from communicable disease

2. Cultures and stocks of agents and associated biologicals Specimens from medical and biological laboratories Cultures and stocks of infectious agents from clinical,

research, and industrial labs

3. Human blood and blood products: these include waster blood, serum, plasma and other blood products

4. Pathological wastes: tissues, organs, body parts, blood and body fluids

5. Contaminated sharps: contaminated hypodermic needles, syringes, scalpel blades, pasteur pipettes, and broken glass

6. Contaminated body parts, animal bedding or animal wastes

7. Miscellaneous contaminated wastes- these include

1. Wastes from surgery or autopsy2. Miscellaneous medical wastes3. Dialysis unit wastes4. Contaminated equipment

Infectious waste management plans Components of the infectious waste management

plan1. Designation of the waste that should be managed as

infectious2. Segregation of infectious wastes from the non-infectious

wastes3. Packaging4. Storing5. Treatment6. Disposal7. Contingency measures for emergency situations8. Staff training

Designation of an infectious waste plan Should specify which wastes are to be

managed as infectious wastes A responsible official or committee should

determine any other miscellaneous materials that should be managed as infectious wastes

Segregating medical wastes

Segregation of infectious wastes at the point of origin

Segregation of infectious wastes with multiple hazards as necessary for management and treatment

Use of distinctive, clearly marked containers or plastic bags for infectious wastes

Use of the universal biological hazard symbol on infectious waste containers as appropriate

Segregating continued

Whenever possible, do not combine medical wastes with hazardous chemical or radioactive wastes

Separate sharps from other medical wastes Sharps should be stored in puncture proof

containers Separate pathology wastes from other

wastes Separate chemotherapy wastes from other

wastes

Mixing wastes

If two types of wastes are mixed, treat mixtures as follows Mixtures of medical and radioactive wastes- decontaminate

the biological components and manage as radioactive wastes

Mixture of medical wastes and hazardous wastes- if safe to do so, decontaminate the biohazardous component and manage as medical waste

Mixtures of medical, radioactive, and chemical hazardous waste- if safe to do so, decontaminate the biohazardous component and manage as radioactive wastes

Packaging of infectious wastes Selecting of packaging materials that are

appropriate for the type of wastes handled: Plastic bags for many types of solid or semisolid infectious

wastes Bottles, flasks or tanks for liquid

Use of packaging that maintains it integrity during storage and transport

Closing the top of each bag by folding or tying as appropriate for the treatment or transport

Place liquid wastes in capped/tightly stopped containers

Handling Sharps

Each year there are some 600,000 incidents of people accidentally struck by needles and sharps

Most common types of risks for spreading blood borne pathogens occurs: Recapping needles Failing to dispose of used needles properly in

puncture resistant containers Accidental breakage of tubes used for collection

of blood of health care settings

Handling Sharps

To protect against needle needle stick injuries, take the following precautions: Avoid needles when safe and effective alternatives are

available Help your employer select and evaluate devices with safety

features that reduce the risk of needle stick injuries Use device with safety features provided by your employer Avoid recapping needles Plan for safe handling and disposal of needles before using

them

Handling sharps

Promptly dispose of used needles in appropriate sharp disposal containers

Report all needle stick and sharp-related injuries promptly to ensure that you receive appropriate follow up care

Tell your owner about any needle stick hazards you observe

Participate in training related to infection prevention

Packaging of sharps

Containers are rigid puncture-resistant containers that, then sealed, are leak resistant and cannot be reopened without great difficulty

Must be red in color, have a biohazard label, be accessible to employees, and be located as close to feasible to the immediate area where sharps are used

Must remain upright throughout use and be replaced routinely. Should not be overfilled to present a hazard

Containers of contaminated sharps will be closed immediately Secondary containers must be closable and prevent leakage

during handling, storage, transport or shipping

Storage

Storage temperature and duration are things to be considered on storage Limiting the storage area near the treatment site Minimizing storage time Proper packaging that ensures containment of

infectious wastes and exclusion of rodents and vermin

Limited access to storage area Prominently displaying the universal biological

hazard symbol on storage area door, waste production etc.

Transport

Avoidance of mechanical loading devices that may rupture packaged wastes

Frequent disinfection of carts used to transport wastes within the facility

Placement of all infectious materials into rigid or semirigid containers before transport off the site

Transport of infectious materials in closed leak proof containers or dumpsters.

Use of appropriate hazardous symbols in accordance with local, state and federal guidelines

Medical Waste Hauling

By a healthcare professional employed by the facility

By contract with a transporter registered with the state

By mail, parcel post, or courier service (sharps only)

Medical Waste Hauling

Transporting by the US. Postal service may be arranged; provided the addressee is a treatment/disposal facility permitted by the state

A mailed parcel of used sharps may be mailed as first class or priority mail. No packages may weigh more then 35 pounds, and the total liquid amount is 50 mls

On-site treatment

Several methods Autoclaving Incineration Thermal inactivation gas/vapor sterilization Chemical disinfection Sterilization by irradiation

Autoclaving

Autoclaving (heating under high pressure) is widely accepted as effective medical waste treatment technology

Steam sterilization, or autoclaving, involves the use of saturated steam within a pressure vessel at temperatures high enough to kill infectious agents in the waste. Sterilization is accomplished primarily steam penetration

Steam sterilization is most effective with low density material such as plastics, metal pans, bottles and flasks.

Autoclaving

• Plastic bags should be placed in a rigid container before steam treatment to prevent spillage and drain clogging.

• Bags should be opened and caps and stoppers should be loosened immediately before they are place in the steam sterilizer.

• Care should be taken to separate infectious wastes from other hazardous wastes.

• Infectious waste that contains noninfectious hazards should not be steam-sterilized.

• Waste that contains antineoplastic drugs, toxic chemicals, or chemicals that would be volatilized by steam should not be steam-sterilized.

Autoclaving

Persons involved in steam sterilizing should be trained in handling techniques to minimize exposure to hazards from these wastes. These should include: Use of protective equipment Minimization of aerosol formation Prevention of spillage during autoclave loading/unloading Prevention of burns from handling hot containers Management of spills

Check autoclave temperature to ensure that the proper temperature is being maintained for a long enough period during the cycle.

Steam sterilizers should be routinely inspected.

Incineration

Newly Regulated Emissions from Medical Waste Incineration:

1. Particulate Matter2. Carbon Monoxide3. Dioxin4. Sulfur Dioxide5. Hydrogen Chloride6. Nitrogen Oxides7. Cadmium8. Lead9. Mercury

Thermal Inactivation

Thermal inactivation involves the treatment of waste with high temperatures to eliminate infectious agents.

This method is usually used for large volumes. Liquid waste is collected in a vessel and heated by heat

exchangers or a steam jacket surround the vessel. The types of pathogens in the waste determine the temperature

and duration of treatment. After treatment, the contents can be discharged into the sanitary

sewer in a manner that complies with State, Federal, and local requirements.

This method requires higher temperatures and longer treatment cycles than steam treatment.

Gas/Vapor Sterilization

Gas/vapor sterilization uses gaseous or vaporized chemicals as the sterilizing agents.

Ethylene oxide is the most commonly used agent, but should be used with caution since it is a suspected human carcinogen.

Because ethylene oxide may be adsorbed on the surface of treated materials, the potential exists for worker exposure when sterilized materials are handled.

Chemical Disinfection

Chemical disinfection is the preferred treatment for liquid infectious wastes.

Consider the following: Type of microorganism Degree of contamination Amount of proteinaceous material present Type of disinfectant Contact time Other relevant factors such as temperature, pH,

mixing requirements, and the biology of the microorganism

Ultimate disposal of chemically treated waste should be in accordance with State and local requirements.

Sterilization by irradiation

Advantages of irradiation: Electricity requirements are nominal. Steam is not required. No heat or chemicals remain the treated waste.

The principal disadvantages are as follows: Capital costs are high. Highly trained personnel are required. Space requirements are great. Worker exposure as a result of leaks in seals or poor work

practices. Disposal of the radiation source may pose problems.

Drain Disposal

Some liquid medical wastes, such as human blood, may be disposed of in sinks if the waste is first autoclaved or brought to a final concentration of 1 percent bleach.

The following medical wastes may not be drain disposed:

Human or animal cultures suspected of containing infectious agents

Cultures and stocks of infectious agents Wastes from the production of infectious bacteria, viruses,

spores, discarded live and attenuated vaccines Non-infectious medical waste disposal should only

be made to sanitary sewers only.

Disposal of treated wastes

Infectious waste that has been effectively treated is no longer biologically hazardous and may be mixed with the disposed of as ordinary solid waste, provided the waste does not pose other hazards that are subject to federal or state regulations.

EPA recommends: Contacting state and local governments to identify

approved disposal options. Discharge of treated liquids and pathological

wastes (after grinding) to the sanitary sewer system. Approval of the local sewer authority must be obtained.

All surfaces, tools, and other objects in contact with potentially infectious materials must be decontaminated and as soon as possible.

Equipment and tools must be cleaned and decontaminated before servicing or being put back to use.

Decontamination should be accomplished by using A solution of 5.25% sodium hypochlorite

(household bleach / Clorox) diluted between 1:10 and 1:100 with water.

Lysol or some other EPA-registered tuberculocidal disinfectant.

Decontamination continued

To clean up spilled blood, you can carefully cover the spill with paper towels or rags, then gently pour your 10% solution of bleach over the towels or rags, and leave it for at least 10 minutes.

To decontaminate equipment or other objects (be it scalpels, microscope slides, broken glass, saw blades, tweezers, mechanical equipment upon which someone has been cut, first aid boxes, or whatever) you should leave your disinfectant in place for at least 10 minutes before continuing the cleaning process.

Cleanup materials must be decontaminated.

Decontamination

Steps to Clean Up a Spill of Animal blood Blood Wear gloves to clean up spill. If broken glass is present, use forceps to pick up

and place in sharps container. Absorb blood with paper towels and discard in

biohazard waste container. Using a detergent solution, clean the spill site of

all visible blood. Wipe the spill site with paper towels soaked in a

disinfectant such as bleach diluted 1:10 Discard all contaminated materials into biohazard

waste container. Wash hands with soap and water.

Questions?