Revising our PHYSIOLOGY practicals :
PLEASE TAKE CARE TO KEEP YOUR
TISSUE PREPARATIONS OR YOUR
PREPARED SLIDES SAFE & INTACT WITH
YOU, LEST ANY LAB. ATTENDANT OR ANY
OF YOUR CLASSMATES THROW IT AWAY
IN THE BIN THINKING IT HAS ALREADY
BEEN CHECKED BY THE EXAMINER !
Why do we stun frogs, instead of using some anaesthetic agent :
The anaesthetic agent of choice (Urethane, or, Di-ethyl Carbamate) is no more available
in the market. We used to prefer this chemical as its use doesn’t depress the CNS, thus
not interfering with our nerve/muscle experiments. Anaesthetic agents like chloroform
and anaesthetic ether etc are potent depressants of CNS, and thus not preferred for our
experiments. Another alternative is pithing of the frog (mechanical destruction of the
brain). Both pithing & stunning are almost equally traumatic for the animal, but stunning
is quite fast, and therefore preferred (of the two lesser evils). Chemicals like Nembutal &
Amytal etc are used for mammals, but are usually not preferred for lower vertebrates like
amphibians etc.
Composition of isotonic saline for frog (amphibian in general)
0.64% to 0.7% NaCl
Composition of isotonic saline for mammals
0.9% NaCl
Composition of amphibian Ringer’s solution
1.0% NaHCO3
1.0% Cacl2
1.0% KCl
0.6% NaCl
001.00 ml x n*
001.00 ml x n*
000.75 ml x n*
097.25 ml x n*
*n = any fixed number
Which of the above solutions do we use for muscle-twitch experiment ?
Amphibian saline
Which of the above solutions do we use for heart perfusion experiment ?
Amphibian Ringer’s solution
Why don’t we use saline solution for heart solution experiment ?
B’coz single ion (cation or anion) solutions (like saline) prove toxic to tissues, whereas a
balanced salt solution (Ringer’s) is NOT toxic. Heart will stop beating soon if we use
only saline to perfuse it.
Why must precavals be ligatured, and bulbous arteriosus left open in the heart
perfusion experiment ?
So that the heart is maximally filled and emptied with each cardiac cycle. If precavals are
not ligatured, Ringer’s fluid will escape through them, thus not filling the heart
maximally – and not letting it obey Starling’s law of heart. Starling’s law : the force of
cardiac muscle contraction is directly proportional to the length of cardiac muscle fibres,
i.e., the more the cardiac muscle fibres are stretched, the stronger (more forcefully) they
contract. So, to make the cardiac muscle fibres stretch maximally, the heart must be filled
up maximally too – and, therefore, we can’t permit any Ringer’s fluid to leak through the
unligatured precavals ! Bulbous areteriosus must be left open to let the Ringer’s fluid
escape post-systole – as naturally happens in the heart in vivo – blood enters through the
precavals & the post caval, and escapes through the bulbous arteriosus.
At what speeds is the drum rotated for muscle & heart experiments ?
640.00 mm/sec (fastest) for muscle twitch
640.00 mm/sec (fastest) for obtaining tuning fork recording
002.50 mm/sec (slowest) for heart
Main precautions for muscle twitch experiment :
1. Take a medium-sized frog (not too small, nor too big). Quick, dry, blunt dissection.
2. As you just begin your dissection, feel the texture of the gastrocnemius, it should be
quite soft & pliable. It should preferably retain that texture & softness till the end. In case
the muscle hardens a bit, you’re not likely to get any good muscle twitchs.
3. The nerve/muscle should not be stretched, overstretched or otherwise damaged, nor
should they be touched with any metallic instruments (it depolarizes the nerve/muscle
membrane). If this is allowed to happen, the muscle may be already tetanized by the time
we are ready to perform the experiment.
4. During the entire dissection, the muscle/nerve MUST be kept moist by constant
pouring of saline solution (not Ringer’s).
5. The circuit should be pre-checked for its operative efficiency. All loose connections
must be tightened, and the drum should be properly leveled, so that the stylus point
touches the smoked paper all the way up and down.
6. Please do ensure that the muscle preparation is rigidly fixed to the wax surface on the
muscle chamber. Please be aware that the pin often comes off.
7. The thread tying the muscle tendon to the stylus must not be loose. Please be aware
that the thread indeed has a tendency to loosen during the experiment.
8. After you have obtained a couple of twitches, please allow the muscle to relax for a
while, and do keep the nerve/muscle moist by constantly pouring saline solution using a
dropper. Please keep your table (and the entire experimental setup) clean and well
organized.
9. As long as you are not ready to obtain twitch recordings, do keep the Du-Bois
Reymond short-circuiting key CLOSED, so that the muscle doesn’t get tetanized by
unnecessary twitching.
10. Don’t begin your experiment with the highest voltage selected from the DC current
source. Increase it stepwise > 2, 4, 6 volts as and when necessary, depending upon the
amplitude of the twitch curve that you desire. Similary, don’t keep the induction coil in
maximum secondary current position – change it as and when required depending upon
the amplitude of the twitch curve that you desire.
11. Don’t forget the hook/weight set in tightening & straightening up the muscle-stylus
writing axis. When you’re not actually recording, just take the hook/weight set off. If
you keep the hook/weight applied constantly, it will unnecessarily put constant load on
the muscle and make it fatigued/tetanised much earlier – rendering it totally useless for
your purpose .
12. Do change the position of the drum (in regard to the metallic axis) once you have
obtained a good twitch, so that your 2nd recording doesn’t overlap and spoil your first
recording.
Main precautions for heart perfusion experiment :
1. Take the biggest possible frog. Quick, dry, blunt dissection – avoiding any major
bleeding.
2. Keep pouring saline on the heart during your dissection.
3. Please pre-adjust the flow of Ringer’s in the perfusion fluid bottle. If necessary, cut the
rubber tubing appropriately short.
4. Please choose the right-sized cannula with a prominent constriction where the post
cavals is to be tied on it.
5. When taking the heart out of the frog’s body, please cut the precavals DISTAL to
(beyond) the ligature points.
6. While obtaining the heart recording please ensure that the Ringer’s solution is getting
properly filled-up in the cannula. Adjust the Ringer’s bottle or the length/direction of the
rubber tubing if necessary.
7. Sometimes you do get better results if you pour back the Ringer’s coming out of the
heart back into the aspirator bottle (duly filtered of course).
8. THE SINGLE MOST CRUCIAL STEP IS : ONCE THE CANNULA IS INSERTED
IN THE POSTCAVAL, LET RINGER’S FLOW FREELY INTO THE HEART, SO
THAT THE HEART IS LITERALLY BLEACHED OF ANY BLOOD – it will start
appearing light pinkish red in color rather than the stark scarlet red color it showed when
the cannula wasn’t inserted. Please conduct your experiment fast (without being hasty &
careless), so that the circulation doesn’t get arrested if the heart isn’t beating, and the
blood doesn’t clot anywhere inside the heart.
9. Once inserted, the cannula MUST NOT slip out of the post-caval opening. If it does, it
may be very difficult reinserting it.
WHETHER YOU HAVE BEEN ASKED OR NOT, MAKE SURE TO CALCULATE
THE LATENT PERIOD, CONTRACTION PERIOD, RELAXATION PERIOD
(TOTAL TWITCH TIME) BY TAKING APPROPRIATE TUNING FORK
RECORDINGS. TUNING FORK RECORDINGS MUST NE TAKEN AT THE
SAME DRUM SPEED (THE HIGHEST) AT WHICH TWITCH RECORDINGS
ARE TO BE OBTAINED.
WHILE OBTAINING ANY RECORDINGS ON THE KYMOGRAPH PLEASE
MAKE SURE THAT THE STYLUS TOUCHES THE DRUM AT A TANGET >
JUST CURVE THE PLASTIC TIP TOWARDS THE DRUM AND MAKE IT
TOUCH THE DRUM IN AN INVERTED ‘L’ SHAPE.
PLEASE MAKE DOUBLY SURE THAT YOU HAVE THE RIGHT DRUM SPEEDS
SELECTED FOR THE EXPERIMENT YOU ARE PERFORMING (see above). YOU
ARE LIKELY TO LEAVE A VERY BAD IMPRESSION IF YOU DON’T CARE
FOR SUCH BASICS.
ONCE MUSCLE-HEART PREPARATIONS ARE TAKEN OUT, DO SPEND
SOMETIME CLEANING THEM. CUT OUT ALL UNNECESSARY TISSUE(S)
ADHERING TO YOUR EXPERIMENTAL PREPARATION WITHOUT
ACTUALLY DAMAGING IT IN ANY MANNER.
YOU DO NEED TO TAKE THE MUSCLE-TWITCH OR HEART RECORDINGS
IN FRONT OF THE TEACHER/EXAMINER. RECORDINGS NOT TAKEN IN
THE PRESENCE OF THE TEACHER/EXAMINER WON’T FETCH YOU ANY
CREDIT. DON’T REMAIN BUSY TAKING TOO MANY RECORDINGS, JUST
ONE/TWO GOOD ONES ARE ENOUGH.
Composition of the fuel that is burnt in the lamp to obtain the soot on the paper :
80 ml of Kerosene oil + 20 ml of used/discarded xylene. Excessive xylene produces very
soft soft, and lower quantities of xylene produce very hard soot. NEVER USE
BENZENE, TOULENE OR CHLOROFORM IN PLACE OF XYLENE. Also, while
sooting, make sure that the paper is exposed just to the soot coming from the flame, and
not to the flame itself – that will almost char the paper and make the soot very hard – you
will see light brown (instead of white) lines wherever the soot is erased !
Composition of the resin that is used to fix (make permanent) the soot on the paper :
Base : Methanol (denature spirit) 100 ml x n*
Additive : Shellac (use in paint/varnish) 25 gms x n*
The mixture is left standing for several days for shellac to dissolve completely, and is
duly filtered before each use.
*n = any fixed number
Why do we conduct muscle/heart experiments on amphibian tissues/organs, and not
on mammalian ones :
Mammalian tissues (warm-blooded vertebrates) have a much higher metabolic rate, and
they are far more susceptible to variations of pH, temperature, O2 concentration,
nutrients etc in the perfusion fluids. Such experiments can definitely be conducted on
mammalian tissues, but doing so requires far more rigorous experimental procedures &
controls. On the contrary, amphibian tissues (cold-blood vertebrates) are far more sturdy,
and do survive a lot longer even during much less rigorous experimental controls &
procedures.
Reagent for preparing haemin crystals : Nippe’s solution (100 mg KBr, KI & KCl in
100 ml of acetic acid).
Reagent for preparing haemochromogen crystals : Takayama solution (a mixture of
pyridine 3ml, saturated solution of glucose 3ml, 10% NaOH 3ml, distilled water 7 ml).
IMPORTANT : We use all three potassium halides in Nippe’s solution as they
hasten the process of color formation in the crystals. This is b’coz KBr, KI & KCl
vary in their photo-sensivity KBr (most) > KI (less) > KCl (least). Haemin crystals
can be prepared using only KCl in Acetic Aid – such a reagent is called
Teichmann’s Solution. The crystals so formed are lightly-colored, and they take
longer to develop whatever little color they do.
PLEASE NOTE THAT THESE BLOOD CRYSTALS ARE JUST BLOODSPECIFIC AND NOT SPECIES-SPECIFIC. AS FORENSIC OR MEDICOLEGAL TESTS THEY DO TELL YOU THAT A GIVEN RED STAIN (SCRAPED
FROM ANY SURFACE) IS BLOOD OR NOT, BUT THEY DON’T TELL YOU
AS TO WHICH SPECIES (OR GENUS FOR THAT MATTER) THEY BELONG
TO – CAT, RAT, HUMAN, FROG ETC.
Reagent for counting RBCs : Hayem’s solution – NaCl 1%, Na2SO4 2.5%, HgCl2
0.25%.
Reagent for DLC & TLC – Dacie’s fluid - 100% acetic acid with a pinch of crystal
violet or Turk’s fluid – acetic acid 3ml, 1% Gentian Violet 1 ml, distilled water 96 ml.
Why do we dilute the blood sample more for RBCs than for WBCs ?
Because RBCs have a far greater numerical presence in our blood than do WBCs. We
dilute our blood 100 or 200 times for RBCs, but only 10-20 times for WBCs.
Reagent(s) for Haemoglobin estimation :
1. 0.1 N HCl (to be taken initially in the reaction tube, till the lowermost graduation).
2. Distilled water (to be used to dilute the blood+HCl mixture for color-matching)
CAUTION : DON’T USE 0.1N HCL for step 2 above !
Please clean the blood-sampling pipettes immediately after use – not to allow any blood
clots to form inside the capillary tube. Also, please pre-check the pipettes for any
previous obstructions in their capillary (just blow air through and feel it coming out on
the back of your hand).
PLEASE memorize comments for the clinical significance of BP apparatus, the Kneejerk hammer for testing patellar reflex, and the Spirometer for respiratory
measurements. While writing comments on sphygmomanometer, please make sure to
add a note on the drug types used to control hypertension. In all these ‘equipment spots’
please comment only upon their clinical significance, and not on the details of the
equipments themselves.
Blood pressure definition : the lateral pressure exerted by blood on the walls of the
blood vessel it is contained in. The smaller the blood vessel, the lesser the pressure and
vice versa.
Starling’s law of heart : see somewhere above !
Please address further queries to/at :
drpkumar@gmail.com
93 501 86363
I will keep updating this page till the 12th of April,
2012.
Best of luck !