Histology - Microtomy

HELLOS.

been experiencing some problems with blogger.. can’t make my postings. but anyway, i’m back to histopathalogy routine lab.. after a month at cytology and a month at processing area of histo lab.. so far what have been shared by the others on histology is staining, processing/trimming, embedding.. i shall touch on microtomy then~ =)

Microtomy, otherwise also known as sectioning, is as we all know from HTECH, the sectioning of tissue blocks using a microtome. After tissues are processed and embedded into paraffin blocks, it is then sent to the microtomy section for sectioning. Each tissue section produced can range from 2-25 microns.

Tissue blocks are first shaved/rough-cut, before it is being sectioned for microscopic viewing. Rough-cutting/Shaving is done at 20 microns per rotation/slice, until the entire surface of the tissue has been exposed. With rough-cutting/shaving, dense and hard tissues which may cause nicks and blunts to the microtome blade can be easily identified, as well as sutures and staples. If tissues are dense and hard, it is will then be pre-treated with RDO (decalcifier) and sectioned last (after all the other tissue blocks have been sectioned), as they might still cause nicks and blunts to the blade. Sutures and staples will be removed if identified. Nicks and blunts on the blade will cause score lines and poor sectioning of the tissue block, which may affect the diagnosis when evaluated by the pathologists.

After blocks are being shaved, it is then placed in 10% fabric softener solution for 5 minutes, and washed in water before placing them on cryoplate for chilling. Chilling of blocks is actually to facilitate fast sectioning, and renders tissue blocks hard for thin sections (normally 3-4 microns) to be produced.

The microtomes are those rotary microtomes, whereby the rotary movement is converted into up and down movement for sectioning, and a forward movement to advance the block for continuous sectioning.

Tissue blocks which are chilled, will be clamped onto the microtome, whereby thickness of the sectioning will be set at 3-4 microns, and section into even thickness and long ribbon of sections. These tissue sections will then be transferred onto the illuminated floatation warm water bath (set at 48 degree Celsius +/- 4 degree Celsius). Tissue sections can also be placed into 1% alcohol floatation bath before transferring to the warm water bath, which serves as an alternative when folds on the tissues are difficult to get rid of. Alcohol have a low vapour pressure which will increase surface tension of the sections, thus aid in the spreading of the tissue sections, so that it can be transferred flat, monolayer onto the microscopic glass slides.

Transferring of the tissue sections onto microscopic glass slides is done by ‘fishing’, whereby the desired sections will be ‘fished’ out of the floatation bath using the glass slides, and subsequently labelled with technican’s (who sectioned the block) initial, accession number (of the specimen), batch number, and block number.

Slides will then be placed on the hot plate (with tissue section facing upwards) for 3 minutes to ensure that the sections will not float off during staining. After which, it will be sent for staining (either H&E by the autostainer, or special stains such as PAS, GMS etc.)

Common faults which may be encountered during sectioning:

1. Ribbons fail to form, which can be due to

• paraffin wax too hard
• blade too blunt
• tilt/angle of blade is too great.

2. Sections compresed, wrinkled or jammed, which can be due to

• Blade too blunt
• Tissue block is still warm
• tile of blade and block is too slight

3. Split ribbon or scratches (score lines) in ribbons, which can be due to

• nicks in the blade
• tilt/angle of blade is too great.
• tissue is too hard
• blade edge is blunt (dull/rounded)

4. Sections full of folds, which can be due to

• floatation bath is too cold/warm
• tissue block is still warm

           Automated Microtome which we use in the lab!

               Controller for the automated microtome (close up)

Cryoplate

Illuminated floatation water bath

Manual Microtome(similar to the one we have in sch!)

Pictures Credit to Lab, taken with permission.

Cheers,

Ang Yu Hui Jacelyn
0702632A

Lab Technique - RT-PCR

RT-PCR stands for real-time polymerase chain reaction. Both PCR and RT-PCR are able to amplify target DNA sequences but the value-addedness of RT-PCR comes from the fact that it can also detect the amplified sequence in real-time. For PCR, agarose gels are required for the detection of the amplified sequence and it can only done after the amplification process. This becomes very time consuming. Because of the increased number of manual steps involved in the detection of PCR products, it is likely that the rate of error due to contamination or other human error is higher than RT-PCR.

(Image taken from http://bio-rad.gene-quantification.info/)

This is a real-time PCR system (CFX96™ Real-Time PCR Detection System from Bio-Rad) that my lab uses. It is able to perform several assays at any one time and is relatively user-friendly.

The procedures to run RT-PCR are easy as the vendor would have provided you with sufficient information to run your samples. For example, for the TaqMan probes that I was using, I was required to add in 10µL of TaqMan Universal PCR Master Mix, 1µL of Assay Mix, desired amount of sample and RNase-free water. The parameters for RT-PCR (not mentioned) and formula are all provided by the vendor. As long as the protocol is followed, every thing should be a-ok.

Lastly, RT-PCR can be used for quantification of gene expression, detection of pathogens and genotyping. There are of course, more applications out there than that that I've listed. It is really how creative anyone can get to exploit this useful machine.

Yvonee Chew 0703189A

P/S Where is Jacelyn?

Bone Marrow Morphology

Hey everyone! Felicia here once again. :)

I'm now attached to the Bone Marrow Morphology Lab for 3 weeks. This lab has the most senior medical technologists and is considered one of the most exciting and challenging labs amongst all the others labs in Haematology. When situated in this lab, one will have the golden opportunity to go down to the wards and experience how the bone marrow is being aspirated out from patients! Indeed it may give one a nauseating feeling, but this is what makes the job in this lab so exciting about. ;)

Bone Marrow Preparation, Staining & Reporting

After clinical history, physical examination and the review of peripheral blood film, bone marrow examination is the next most important step used in the diagnosis of haematological disorders. It is the only way used in the classification of leukaemias and myelodysplasias. It allows the assessment of the body iron stores, marrow cellularity, myeloid to erythroid ratio, maturation of cell lines and relative quantitation of eosinophils, lymphocytes and plasma cells. In addition, infiltration by fibrosis and other malignant cells can also be diagnosed. It is also used to assess response to treatment in leukaemias and to assess prognosis in aplastic anaemia and agranulocytosis. The absence of iron stores help to differentiate iron deficiency anaemia from other hypochromic anaemias. Trephine (a fixative for the bone) biopsy may help to diagnose myelofibrosis, aplasitc anaemia, malignant lymphoma or secondary carcinoma.

Equipments & Materials

1. Microscope
2. Multitimer
3. Differential counter
4. Glass slides
5. Spreader
6. May-Grunwald stain
7. Giemsa stain
8. Buffered distilled water pH8.6
9. Mounting medium DPX
10. Adhesive labels
11. Reporting worksheets

Specimen Collection

The marrow aspirate can be obtained from the posterior superior iliac spine due to its accessibility and relative safety which a sample can be obtained.

Preparation of bone marrow smears

Bone marrow smears can be made directly at the beside during aspiration procedure. The sample is then sent to the lab for slide preparation.

Wedge smears
About 0.3ml of bone marrow from the first stringe is put onto a clean glass slide to allow the blood to drain away onto another slide. The marrow fragments tend to adhere to the slide and most of them will be left behind. A smooth edge spreader of not more than 2cm in width is used to make 10-12 wedge smears of size 2x3cm. The marrow fragments are then dragged behind the spreader and they leave a trail of cells behind them. Differential counts are then performed by the senior staff.

Squashed preparation
A few particles are placed on the centre of a clean glass slide in which another slide is gently compressed to spread and disperse the particles as the slides are pulled apart.

Trephine imprint
Several touch or imprint of the bone fragment are prepared by gently touching the core and rolling between 2 slides.

Handling conditions

1. All bone marrow smears are labeled with the patients' initials immediately after preparation. They are kept in separate trays and brought back to the lab with request forms to be filled in by staff.

2. A unique marrow number is assigned to each patient sample and are written in request forms, record book etc.

3. 2 wedge smears, 1 blood film, 1 squashed preparation and an imprint are selected for May Grunwald Giemsa staining. Another wedge smear with visible particles is used for iron staining. Controls with normal/increased iron stores are stained in parallel with the patients' smear.

Bone Marrow Smear - May Giemsa Stain (x1000)

Image from: pathy.med.nagoya-u.ac.jp/atlas/doc/node76.html

4. A set of buff cards and adhesive labels for storing the smears together with a worksheet with patient's details are prepared.

5. Any unstained smears are kept in closed containers for 2 weeks for further tests, if necessary.

I've only spent 2 days in this lab, so this is whatever I've observed and learnt so far! If you guys have any queries, please do not hesitate to ask. ;)


Signing off,
Felicia
0703345I

HELLO PEOPLE! QINGLING'S BACK! :D

This month, I'm posted to the Thalassaemia section of the lab! The one that Felicia was at before me! I've learnt to perform five tests there and surprisingly it was pretty easy! My colleagues in the same lab as me were awesome! They taught me every test patiently despite me making mistakes along the way! They even left the test for me to perform all by myself whenever the test was ordered! So before i begin, im sure all of us know what a sickle cell looks like right? yes. like this!

Picture taken from

http://repairstemcell.files.wordpress.com/2009/03/sicklecell.jpg

Therefore, in this entry, I’m going to share with you the Sickling Test! It provides a rapid screening test for Hb-S status of an individual and serves as a confirmatory test for the Alkaline Cellulose Acetate (the one that Felicia mentioned in one of the previous post) and Acid Gel electrophoresis. One important characteristic

of this test is that the sickling phenomenon occurs only with low oxygen tension. It will detect if the RBC will change into a sickle shape after mixing with the chemical that will

reduce the amount of oxygen carrying it.

A positive and negative control should be used in this test.

The method is as follows.

First, we have to prepare the reagent, which is a 0.2g of Sodium Metabisulphite. It is measured accurately using the weighing balance. After which, dissolve with 10mL of deionised water and mix it up and down using a pipette. 5 drops of the reagent is then added to 1 drop of anti-coagulated blood on the slide and mixed well. Immediately cover it with a cover-glass and leave for it to dry completely as the mixture may be present outside the cover glass. Seal the sides of the cover-glass with petroleum jelly-parrafin

wax mixture. Ensure that it covers the sides of the cover-glass properly as no oxygen should enter. Incubate at 37C for 2 hours. Finally, examine under microscope.

Sickling test should be negative in a normal individual. However in patients with Hb-S disease,

a marked sickling is usually visible.

One important thing to note while performing this test is that patients on medication of drugs such as phenothiazines dapsone will give a false negative result. Also old reagents or if preparation

is improperly sealed, will also result in false negative result.

- - -

Isn't it simple? :D

If you have any doubts, dont hesitate to ask me!

Immuoprecipitation (IP) using Dynabeads

(Taken from http://images.google.com.sg/imgres?imgurl=http://www.invitrogen.com/etc/medialib/en/images/mainbody/data/image.Par.55626.Image.-1.0.1.gif&imgrefurl=http://www.invitrogen.com/site/us/en/home/brands/Dynal/Magnets.html&usg=__YukqNWjNDOfL_tHRNjAEQyUhOaM=&h=163&w=180&sz=7&hl=en&start=4&tbnid=eqmFkGj0Kb2_cM:&tbnh=91&tbnw=101&prev=/images%3Fq%3Ddynamag%2Bspin%26gbv%3D2%26hl%3Den)

Immunoprecipitation (IP) using Dynabeads are more efficient and reliable.
It has several advantages over the Crosslink Immunoprecipitation method.

• Reduce protocol time as short to 30 minutes only instead of a 2 day experiment
  No need for time-consuming procedures e.g. preclearing of cell lysates, 1 minute spin down each time etc.
• Materials will not be lost from spun-down resins or excess surface during pipetting
• Reduce the comsumption of expensive antibodies
• Fast and gentle method that causes minimal physical stress to target proteins
• Able to work with concentrated proteins solutions; therefore dilution is not required
  

Binding of Antibody (Ab)

1. Completely resuspend Dynabeads by pipetting or rotating on a roller (5 min).
2. Transfer 50 µl Dynabeads to a tube, place on magnet and remove supernatant.
3. Remove tube from magnet and resuspend the Dynabeads in 200 µl Ab Binding &
   Washing Buffer containing your Ab of choice. (Typically 1 - 10 µg Ab, the optimal
   amount needed will depend on the individual Ab used).
4. Incubate 10 minutes with rotation at room temperature.
5. Place tube on magnet and remove supernatant.
6. Remove tube from magnet and wash the Dynabeads-Ab complex by resuspending
   in 200 µl Ab Binding & Washing Buffer.

Immunoprecipitation of Antigen (Ag)

1. Place tube on magnet and remove supernatant
2. Add your Ag-containing sample (typically 100 - 1,000 µl) to the Dynabeads-Ab
   complex and gently resuspend by pipetting.
3. Incubate 10 minutes at room temperature with rotation.
   Place tube on magnet, transfer supernatant to a clean tube.
4. Wash the Dynabeads-Ab-Ag complex 3 times, using 200 µl Washing Buffer for
   each wash. Mix gently by pipetting.
5. Resuspend the Dynabeads-Ab-Ag complex in 100 µl Washing Buffer and transfer
   the suspension to a clean tube. Place tube on magnet and remove supernatant.

Elution of Ab/Ag complex (alternatives A: denaturing or B: non-denaturing)

• A. Gently resuspend the Dynabeads-Ab-Ag complex in 20 µl Elution Buffer. Add 10 µl
  NuPAGE® LDS Sample Buffer / NuPAGE® Reducing Agent mix and incubate 10 minutes
  at 70°C. Place tube on magnet and load supernatant/sample onto a gel. (Alternatively,
  the Dynabeads-Ab-Ag complex can be resuspended in the SDS sample buffer of your
  choice and heated as per your standard protocol prior to gel loading.)
• B. Gently resuspend the Dynabeads-Ab-Ag complex in 20 µl Elution Buffer. Incubate
  2 minutes at room-temperature. Place tube on magnet and transfer supernatant/sample
  to a clean tube.

However although the kit is fast and easy to use, however the yield is relatively little and there is alot of "background noise" due to the presence of heavy and light chains from antibodies (co-elution due to the lack of crosslinking in this protocol, therefore there is antibody contamination).
Background noise - Antibody contamination that leads to the production of a heavy and light chain bands appearing on the blot, hence rendering it hard to determine which band is the protein band i.e. immunoprecipitated.

I am currently trying to optimize this protocol by trying out preclearing of lysate (prevent nonspecific binding), incubating with sample and antibody for a longer period of time (to enhance yield) and crosslinking using DSS to prevent co-elution of antibodies; so as to produce better results in future.

Li Yinliang Alex
TG02 0704894E
Group 8
3 August 2009

Lab Technique - Electrochemistry

Here's something really different for all of you out there. My internship focuses mainly on electrochemistry which is the study of chemical reactions involving the transfer of electrons between electrodes and electrolytes.

(Picture taken from http://www.als-japan.com/1031.html)

Electrodes come in different sizes and differ in the inert material in the middle. This picture shows a few glassy carbon electrodes. For a gold electrode, the middle will be gold instead of grey.

Electrodes are electrical conductors that can be used for various purposes such as electrocardiography (ECG) and chemical analysis using electrochemical methods. There are different types of electrodes available in the market but the ones that I have been using are the gold (Au) electrodes.

In order to ensure that the Au electrodes are fit for use, they are polished to remove unwanted chemical residues from previous use. New electrodes of course, do not need to be polished. Polishing is done using alumina powder on a nylon pad. The electrodes are held perpendicular to the nylon pad (with Au in contact with the pad that has alumina powder.) Then for a desired time (eg 10 min each), these electrodes are polished in a figure of 8 or circular motion. The particles in the alumina powder will remove the residues on the gold surface. In addition, nano-strip can be used too. By incubating the electrodes in nano-strip for 15 min, any remaining organic residues are removed.

So how do you know if the electrodes are clean? Electrochemical signals are measured with an electrochemical workstation. A clean Au electrode should give a high electrochemical signal while a Au electrode covered with residues will give a low signal because these residues will slow down the transfer of electrons between the electrode and electrolyte.

(Picture taken from http://www.uni-muenster.de/Physik.PI/DeCola/equipment.html)

This is a picture of an electrochemical workstation. The electrodes are connected to the workstation with the use of wires so that a small amount of voltage can be passed through.

Yvonee Chew 0703189A

Cytology - Processing Gynaecological Specimens

Heys!

it's already end of week 4 of SIP. 16 weeks more to go. 4 mths more to go!

anyway. i'm attached to the histopathology/cytology department. over at the hospital i'm attached to, histopathlogy actually sort of belong to the same department, though both deals with different type of specimens, but is usually inter-related.

we didn't get to do much during our first week, even though for that whole week we were at histo department. from week 2 onwards, we are all allocated at different sections. as for me, i'm now at cytology lab for 1mth, so for this post, i'll be talking about things that are done at our cytology lab. =)


Cytology is actually the study of cells to aid in the diagnosis of diseases. In our lab, we deal with 2 different groups of specimen, Gynaecological (gynae) and Non-gynaecological (NG) specimens. Gynae specimens are actually just cervical pap smears, while NG specimens refers to body fluids (peritoneal, pleural, pericardial, esophageal washings, bronchial washings/aspirates etc), CSF, sputum, urine, and fine needle aspirations (knee, breast, thyroid, bones etc). I'm now covering both gynae section and NG section, but due to safety reasons, we're actually not allowed to perform hands-on for NG specimens, so yeah, all we can do is observe and learn the techniques. =) I'll cover on the gynae section because that's what i've been doing most of the time.

Gynae specimens usually comes in two different type. Liquid-Based preparation or Conventional Smears.

SIP slides on CYTO - gynae

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=))


Ang Yu Hui
0702632A