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Article 9 course > Administration & Extraction Methodology > Flashcards

Administration & Extraction Methodology Flashcards

1
Q

What is covered regarding Experimental procedures
In laboratory animal science?

A

Administration of drugs
Collection of blood
Collection of urine and feces

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2
Q

In what two broad ways can a drug be administered (application & effect)

A

Systemic & Local

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3
Q

What is meant by systemic?

A

Reaches the circulatory system so the entire body is affected

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4
Q

What is the entry route for local application?

A

‘Topical’

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5
Q

What two entry routes are associated with systemic application?

A

Enteral administration
- Via the Gastrointestinal tract
Parenteral administration
- Everything that is not enteral

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6
Q

Give two examples of topical application

A
  • Cutaneous (on the skin)
  • Mucous membrane
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7
Q

Give 5 examples of mucous membranes that can be used to topical application

A

eyes
ears
nose
lungs
vagina

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8
Q

Give 3 examples of enteral application

A

Injection
Respiratory
Cutaneous (?)

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9
Q

Give two examples of parenteral application

A

Oral
Rectal

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10
Q

Give three stages to consider for deciding on a specific drug delivery method

A

Administration/ Absorption
Distribution/ metabolism
Elimination

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11
Q

Describe the route when a drug is administered intravenously

A

Intravenous administration -> blood
Eliminated via kidney (urine) and skin (sweat)

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12
Q

Describe the route when a drug is administered orally

A

Oral administration-> gut -> liver via portal system -> blood
Eliminated via gut (feces), kidney (urine) and skin (sweat)

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13
Q

Describe the route when a drug is administered subcutaneously

A

Subcutaneous administration -> blood
Eliminated via kidney (urine) and skin (sweat)

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14
Q

Describe the route when a drug is administered transdermally

A

Transdermal administration -> blood
Eliminated via kidney (urine) and skin (sweat)

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15
Q

Describe the route when a drug is administered transdermally

A

Inhalation -> Lungs -> Exhaled air

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16
Q

Give 8 things to consider when deciding a specific drug method

A
  • Local or systemic action
  • Site of desired action
  • Physical and chemical properties of the drug
  • Rapidity of response desired
  • Extend of drug absorption
  • Effect of digestion and first pass metabolism
  • Accuracy of dosage required
  • Condition of patient/animal
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17
Q

Is an aseptic technique required for:
Enteral administration?
Parenteral administration?

A

Enteral administration: Not required
Parenteral administration: Required

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18
Q

What is meant by the first pass effect?

A

Hepatic metabolism of a pharmacological agent; Metabolic process of the liver will mean that some of it will be dissolved and passed through the urine, some are deactivated by the liver, some are activated. Generally the greater the first pass effect, the less will reach the systemic
circulation.

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19
Q

Is there a first pass effect in enteral administration?

A

Oral: Yes
Rectal: No

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20
Q

How is rectal administrated carried out in small lab animals?

A

It is not

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21
Q

How may a drug be orally administered?

A

● Drinking water
● Oral cavage

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22
Q

What are the benefits of placing it in food or drinking water?

A
  • Easy to do
  • No disturbance of the animal
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23
Q

What are four attention points of placing it in food or drinking water?

A
  • Variation in consumption
  • Substance solubility
  • Substance stability
  • Substance smell/taste
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24
Q

Why is there a variation in consumption with oral adminitration?

A

Lack of control over how much animal takes in; can weigh a bottle as an indication but no exact way of telling.

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25
Q

Why should you be concerned with the taste or smell of a solution with oral administration?

A

If a substance has a very sour taste for example they will not want it, can be used as a water restriction technique.

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26
Q

What is oral cavage/ stomach tube?

A

Passage of a gavage needle into the esophagus, and this technique often involves the animal swallowing the gavage needle as it approaches the pharynx.

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27
Q

What is the benefit of using a oral cavage/ stomach tube?

A

Direct accurate individual dose, no variation in
consumption

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28
Q

What are the attention points of using an oral cavage technique?

A
  • Disturbance of the animal
  • Still first pass effect
  • Practice is required for restraining the animal and introducing the needle into the oesophagus and not the trachea
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29
Q

What points of attention does the oral cavage neutralise?

A

– Substance stability
– Substance smell/taste
– Variation in consumption

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30
Q

How can the stress during an oral cavage be attenuated?

A

Habituation

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31
Q

Describe the steps involved in the oral cavage technique

A
  1. fill the syringe and attach the cannula
  2. Flexible cannula are preferred as they are less likely to cause damage to the oesophagus
  3. Excess agent should be wiped from the cannula to avoid introducing unpalatable material into the mouth
  4. The length of cannula required to reach the stomach is measured on the outside of the animal
  5. Mouse should be restrained by the scruff sufficiently firmly so that it is not able to move its head during the procedure
  6. If restraint is too firm, this can result in vocalisation and other signs of distress
  7. Tube is passed down animals mouth an into stomach, should be no or minimal resistance encountered when passing the tube.
    - Important that the head and oesophagus are straight so you have clear passage through
  8. Resistance could indicate insertion into the trachea, then the tube should be withdrawn and re-inserted
  9. Tube should be administered slowly to minimise reflux from the stomach
  10. After dosing is complete, tube should be gently withdrawn, animal carefully returned to cage and observed to ensure no immediate adverse effects. If signs of respiratory distress are seen, animal should immediately be humanely killed.
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31
Q

How is rectal administrated carried out in small lab animals?

A

It is not

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32
Q

How is rectal administrated carried out in small lab animals?

A

It is not

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32
Q

Why is length an important factor for the cannula used in an oral cavage?

A

Too short might not reach stomach, too long and it can be dangerous for the animal

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33
Q

How is rectal administrated carried out in small lab animals?

A

It is not

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33
Q

Why may a flexible cannula not be preferred?

A

The flexible tube carries risk that if the animal is not well restrained they can bite through it in one bite and the tube may become stuck.

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34
Q

Why is aseptic adminitration required in parenteral administration?

A

No degredation by GI tract; most materials will not survive acidity of stomach

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34
Q

How is rectal administrated carried out in small lab animals?

A

It is not

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35
Q

Name four forms of injection

A

Subcutaneous
Intra-muscular
Intra-peritoneal
Intra-venous

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36
Q

Give 9 basic rules of injections

A
  • Pick correct needle size
  • Use a clean, sharp, sterile needle
  • Fill syringe free of air bubbles
  • Fluids should be room or body temperature
  • Restrain the animal
  • Inject with beveled (open) side of the needle upward
  • Aspirate to validate the injection site
    • Fluid entering your syringe means you have to reposition!
    • Vein/bladder/intestine
  • Inject slowly
  • Don’t inject more fluid than the recommended maximum volume
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37
Q

What does the size of the needle depend on?

A

Size depends on animal and location

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38
Q

How many times is it safe to use a single needle?

A

Once

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39
Q

What should you do if taking a solution from a fridge?

A

Warm it first

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40
Q

What does it mean to aspirate for verification?

A

Can check if you are in a vacuum to verify, although some say you can create a vacuum with the needle and taking up tissue can be dangerous.

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41
Q

Why should you inject slowly?

A

Injecting too fast can exert force and do damage, although too slowly can be stressful for you & animal.

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42
Q

What is the recommended maximum volume?

A

Depends on species and injection method

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43
Q

For a mouse give:
The recommended dose (when the animal displays discomfort) in ml/kg
for:
oral
s.c
i.p.
i.m.
i.v. (bolus)
i.v. (slow inj.)

A

oral: 10 (50)
s.c: 10 (40)
i.p.: 20 (80)
i.m.: 0.05 (0.1)
i.v. (bolus): 5
i.v. (slow inj.): 25

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44
Q

For a rat give:
The recommended dose (when the animal displays discomfort) in ml/kg
for:
oral
s.c
i.p.
i.m.
i.v. (bolus)
i.v. (slow inj.)

A

oral: 10 (40)
s.c: 5 (10)
i.p.: 10 (20)
i.m.: 0.1 (0.2)
i.v. (bolus): 5
i.v. (slow inj.): 20

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45
Q

For a macaque give:
The recommended dose (when the animal displays discomfort) in ml/kg
for:
oral
s.c
i.p.
i.m.
i.v. (bolus)
i.v. (slow inj.)

A

oral: 5 (15)
s.c: 2 (5)
i.p.: * (10)
i.m.: 0.25 (0.5)
i.v. (bolus): 2
i.v. (slow inj.): *

  • Data not available
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46
Q

What should be taken into account for multiple injections?

A

Take maximum fluid dose per
minimum time past into account. Approximately 10 ml/kg per 30min.

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47
Q

How should you interpret these injection volume values?

A

This is a high value and you would not want to reach it. New study shows new lower maximum volumes; Talk to animal staff or relevant people for the max volume in your institution.

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48
Q

Describe a subcutaneous injection

A

Subcutaneous injection is used to deliver a liquid substance under
the skin.

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49
Q

Describe an intramuscular injection

A

Intramuscular injection is used to deliver small amounts of liquid substances to the animal via the muscle

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50
Q

Describe an intraperitoneal injection

A

IP injection is used to deliver larger amounts of liquid substances to the animal

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51
Q

Compare the uptake of s.c, i.m, i.p and i.v

A

s.c: Relatively slow absorption (few bloodvessels in SC tissue); slow elongated onset

i.m: Quick uptake (many bloodvessels in muscular tissue); quick onset

i.p: Quick uptake through the vasculature of the peritoneum

i.v: Fast uptake, gotten rid of quicker

52
Q

Compare the volumes of s.c, i.m, i.p and i.v

A

s.c: Volumes are higher than other methods, (which may play into your decision; 40 is very high and rarely used in practice.)

i.m: Small volumes to prevent
pain and tissue/nerve damage

i.p: Relative large injection volume

i.v: Small volumes

53
Q

Compare the discomfort of s.c, i.m, i.p and i.v

A

s.c: Can be irritating

i.m: Local pain (Tissue is quite dense + sensitive and can be painful, less volume possible as less space.)

i.p: *

i.v: *

54
Q

Why may s.c injections be more flexible than other methods?

A

Flexible in that if multiple injections are possible (e.g scruff or flank- in front of hind leg) within an animal, multiple sites are possible.

55
Q

Describe the process of a s.c injection

A
  • Use the neck scruff restraining method
  • The needle is placed into the loose skin of scruff of the neck in mice or rats
  • The needle slides between your index finger and thumb

Max volumes:
– Mouse 10 - 40 ml/kg total; divided between 2-3 sites
– Rat 5 - 10 ml/kg total; divided between 2-4 sites

56
Q

Describe the process of an i.m injection

A
  • multiple restraining methods possible (e.g shoulder & tail restraint with assistant)
  • Inject caudally of the sciatic nerve
    This nerve runs along the length
    of the femur (front or back)
  • Important to avoid sciatic nerve
  • Important to consider the angle: to go deeper, avoid the nerve etc, can feel the muscle of a given animal to get an idea of this.

Max volumes:
– Mouse 0,05 - 0,1 ml/kg/site; max 2-4 sites
– Rat 0,1 - 0,2 ml/kg/site; max 2-4 sites

57
Q

Why is a high concentration necessary for i.m injection?

A

need to have high concentration as very limited by volume.

58
Q

Describe the process of an i.p injection

A
  • Use a short thin needle
  • Take care to avoid internal organs
    • If you go too deep you can inject into intestine instead of the cavity
  • Inject next to the mid-line to avoid the bladder
  • Use an appropriate injection angle
    • Too horizontally → subcutaneous
    • Too vertically → intestines
  • Aspirate to validate injection site
    • If you see brown or yellow substance then you are not in the appropriate location and will have to redo the injection.
59
Q

What is sometimes recommended when injecting i.p but is contested by research? (apart from aspiration)

A

Leaning the mouse forward may shift intestines forward to avoid mistaken injection but other research shows this doesn’t make a difference (except maybe for more experienced researchers.)

60
Q

How can the internal organs of the animal be damges when injecting i.p? What can you do when one of these sites are hit?

A

Damage is minimal to the intestines or bladder until you begin injection of solution. If one of these sites are hit, it may be best to switch to the other side of the animal.

61
Q

Where are i.v injections typically in mice, rats and rabbits?

A

Tail vein in mice and rats
Ear vein in rabbits

62
Q

How are rodents often restrained when injected i.v?

A

A restraining device (tube) is often used

63
Q

Why is it important to be stable when injecting i.v?

A

even loose movements may cause you to leave the vein

64
Q

How should you angle when injecting i.v?

A

should enter at an angle to go along with the vein almost parallel

65
Q

Where in the tail should you inject?

A

Better higher in the tail, lower it can be harder to enter the vein. If you miss you can only re-enter higher at the tail, so better not start at the base.

66
Q

How many tries should you get with injecting i.v in the tail?

A

Vessels also have a habit of contracting once you damage them and thus you only have limited (2-3) tries.

67
Q

What is important after injecting i.v? Why? (2)

A

Important to place pressure following injection to promote blood clotting and prevent the injected solution from flowing out.

68
Q

When else may a restraining tube be used?

A

Also specialised research where they place animal tube to restrain them and drug is in air circulation, e.g for respiratory research or covid.

69
Q

Name two other specialised techniques for targeting a specific organ

A

Brain → stereotactic surgery
Respiratory – Inhalation anesthesia
– Termination method (CO2)

70
Q

What two forms of cutaneous parenteral admistration are there?

A

Epidermal & transdermal

71
Q

Compare epidermal and transdermal techniques in regards to their application & effect

A

Epidermal
– Stays in the epidermal layer
– Acts local

Transdermal
– Delivered across all layers to reach the systemic circulation
– Acts systemic

72
Q

How may cutaneous methods be administered? (4)

A

Drops
Sprays
Ointments
Transdermal patches

73
Q

What two transdermal pathways are there?

A
  • Transcellular
  • Intercellular
74
Q

What is transdermal delivery highly dependant on?

A

Transdermal delivery is highly dependent on the physical and chemical properties of the drug

75
Q

Do cutaneous delivery methods have a first pass effect?

A

No

76
Q

How stable are cutaneous methods and are they long lasting?

A

Administration can be stable and long lasting (transdermal patch)

77
Q

Comment on the onset of cutaneous methods

A

Slow onset

78
Q

Comment on the accuracy of cutaneous methods

A

Often not very accurate

79
Q

Comment on the discomfort of cutaneous methods

A

Substances may be irritating to the skin/mucous tissue

80
Q

What may impede delivery of cutaneous methods specific to animals?

A

Animal may lick off the drug

81
Q

Comment on the application and effect of topical administration

A

Local application and effect; applied cutaneously (epidermic)

82
Q

Give three reasons why you may want to take blood samples from lab animals

A
  • For analysis of biochemical, metabolic, toxicological or immunological parameters.
  • For examination or culture of micro-organisms.
  • For production of antibodies.
83
Q

What may your choice of blood collection technique depend on? (8)

A
  • The purpose of the blood collection, e.g. biochemical analysis, DNA extraction etc.
  • The minimum volume required for analysis
  • The duration and frequency of sampling.
  • The impact on animal welfare.
  • The need for aseptic obtained blood
  • The need for an arterial versus venous sample.
  • The suitability of sedation and/or anesthesia
  • The potential for stress-induced effects on biochemical and haematological parameters.
    • stress can induce certain biomarkers
      in the blood.
84
Q

Give four general principles of blood sampling

A
  • Volume and number of samples should be kept to a minimum.
  • Maximum volume depends on the total blood volume of the animal.
  • Take account of the combined effect of sample volume and the frequency of sampling.
  • Data interpretation and scientific validity may be confounded if excessive sampling is employed.
    • can impact animal health and in consequence your experiment.
85
Q

How much blood do mice have on average?

A

On average, mice have around 72 ml/kg blood.

A mouse weighing 25 g has approximately 1.8 ml total blood volume.

86
Q

How much blood do rats have on average?

A

On average, rats have around 64 ml/kg blood.

A rat weighing 300 g has approximately 19.2 ml total blood volume.

87
Q

What are the consequences of rapid blood removal and how much is safe?

A
  • 10% TBV can be safely removed on a single occasion
  • With fluid replacement (e.g saline) up to 15%
  • Most animals go into shock if 25-30% TBV is rapidly removed
  • Over 50% die if 30-40% TBV is removed
  • Nearly all die if more than 40% TBV is removed
88
Q

How are replacement fluids administered?

A

Replacement fluids can be given subcutaneously and should be warmed

89
Q

What are the consequences of (safe) blood removal?

A

The issue is not fluid loss; blood volume is rapidly replaced (24h). Other blood components may not be restored for several weeks
(Hematocrit, hemoglobin, etc. )

90
Q

How much blood can thus be removed in practice?

A

A maximum of 1.0% TBV can be removed every 24 hours

91
Q

In a single sampling case (toxicity study), what is the approximate recovery period of 7.5%, 10%, 15% circulatory blood volume removed?

A

7.5%: 1 week
10%: 2 weeks
15%: 4 weeks

92
Q

In a multiple sampling case (toxokinetic study), what is the approximate recovery period of 7.5%, 10-15%, 20% circulatory blood volume removed?

A

7.5%: 1 week
10-15%: 2 weeks
20%: 3 weeks

93
Q

Give two attention points for blood collection

A

Check mice for signs of distress or anemia (rapid breathing, pale color of mucous membrane, muscle
weakness).

Check mice for local trauma, infection or irritation at collection site.

94
Q

Give four vials for blood collection

A
  • Heparin or EDTA tubes
  • Eppendorf tubes
  • Capillaries
  • Vacuum tubes
95
Q

Give 6 sites for extracting blood in rodents

A

Tail vein
Saphenous vein
Dorsal pedal vein
Ear vein in rabbits
Orbita punction
Facial vein

96
Q

Describe the process of extracting blood from the tail vein

A
  1. Place animal in restrainer
  2. Warm tail to dilate vessels
  3. A small shallow incision can be made on the tail to collect the blood
  4. A needle can be placed in the vein to collect the blood
  5. After withdrawal put slight pressure on the tail
97
Q

Describe the process of extracting blood from the facial vein

A
  1. The facial vein (superficial temporal vein) can be punctured in awake mice
  2. When the neck scuff is held firmly the blood will flow easily
  3. When you loosen the grip on the neck scruff the bleeding will slow or stop
98
Q

What is a benefit of the facial vein?

A

Less risk and less invasive than orbital puncture!

99
Q

When is topical administration typically used in animal research?

A

Not very common for research apart from eye ointment to prevent dehydration of the eyes.

100
Q

What is worth noting about herapin and EDTA tubes?

A

Heparin influences PCR, might want to use EDTA, some tests are the opposite. Both would not be suitable for analysing plasma as you want the blood to coagulate and these inhibit clotting

101
Q

What are the benefits of using the facial vein?

A

Easier vein to identify and puncture and less risk and less invasive than orbita puncture!

102
Q

What are the negatives to using the facial vein?

A

It can be harder to control the blood blow while restraining.

How much you can collect is limited.

Cannot make multiple injections .

103
Q

What is the purpose of using a glass capillary and what is a downside?

A

Glass capillary allows blood to flow into it without puncturing the vein; cannot do this repeatedly.

104
Q

When is an orbita puncture typically used?

A

Used on small rodents
Under anesthesia

105
Q

What is typically used to carry out an orbita puncture?

A

Small glass capillary into the orbital vessels (Retro orbital sinus via medial canthus)

106
Q

Is a saphenous vein puncture typically used in rats and mice?

A

Yes

107
Q

Where is the saphenous vein?

A

Runs along the femur

108
Q

Is anaesthesia necessary for a saphenous vein puncture?

A

No

109
Q

Where is the dorsal pedal vein?

A

Dorsal sole of paw

110
Q

How much blood is drawn during exsanguination?

A

Obtain the maximum amount of blood (Even more than cardiac; up to 50% of total blood volume)

111
Q

How is exsanguination carried out in lab animals?

A
  • Decapitation
  • Puncturing the aorta
112
Q

What else is required of the researcher to identify the sapheus vein?

A

Shaving of the leg

113
Q

How much blood can be drawn from the dorsal pedal vein & is anaesthesia required?

A

Small amount & no anaesthesia required

114
Q

When is the marginal ear vein typically used?

A

The marginal ear vein is commonly
used to collect blood from rabbits

115
Q

Comment on the accessibility of the ear vein and the requirement of anaesthesia

A

The vein is easily accessible and visible
No anaesthesia needed

116
Q

How much blood can be drawn from a cardiac puncture?

A

Cardiac puncture is used to obtain large amounts of blood; Blood collected directly from the heart

117
Q

When is a cardiac puncture typically used?

A

Always under anaesthesia

Only appropriate for terminal experiment

118
Q

How can daily blood sampling in a painless and stressless manner be carried out?

A

Cannulation; technique in which a cannula is placed inside a vein to provide venous access. One invasive procedure and after bloodflow can be controlled for painless and stressless sampling to take place

119
Q

What is a downside to cannulation?

A

Blood clotting in cannula, maintenance is required (flushing with anticoagulant)

120
Q

Is aseptic exsanguination possible?

A

No, it is not sterile

121
Q

Is aseptic cannulation possible

A

Yes

122
Q

To recap, what are the recommendations if you require more than one blood sample from the same mouse in terms of blood volume extracted?

A

Maximum <10% TBV on a single occasion AND <15% TBV in 28 days.

For repeat bleeds at short intervals, suggested limit <1% TBV in 24 hours AND consider cannulation

123
Q

To recap, what are the recommendations if you one blood sample from the same mouse in terms of blood volume extracted?

A

Maximum <10% TBV

OR terminal sample under general anesthesia (volume unrestricted)

124
Q

If you require less than 0.2 ml of blood what options do you have where general anaesthesia is not required? (6)

A

Saphenous vein
Dorsal pedal vein
Tail vein (mouse)
Facial vein
Bloodvessel if cannulated
Marginal ear vein (rabbit)

125
Q

If you require less than 0.2 ml of blood what options do you have where general anaesthesia is required? (4)

A

Sublingual vein (drains from the tongue)
Retro-orbital
Jugular vein (rat only)
Tail clip

126
Q

If you require more than 0.2 ml of blood what options do you have?

A

Non-recovery methods:
Cardiac puncture
retro-orbital
Decapitation

127
Q

What is typically used for the collection of both urine and feces?

A

Metabolic cage

128
Q

Describe the concept of a metabolic cage

A

Animal placed on grid for length of time, Food and water is controlled and urine and feces is separated as it is secreted allowing for quantitative collection.

129
Q

What if only a few droppings are needed?

A

When only a couple of droppings are needed you can put a mouse or rat in a clean cage for a couple of minutes

130
Q

What can be used only for the collection of urine?

A

Catheterisation of the bladder

131
Q

How can you promote rodent urination without the introduction of extra fluids?

A

Rodents usually urinate when you place
them on a cold surface

132
Q

What other body fluids are also often extracted?

A

Cerebrospinal fluid (CSF)

Bile → ductus choledochus

133
Q

How can CSF be collected?

A

Cannula between skull and cerebellum, tap above the head then allows to multiple collections of CSF.

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