Lab Techniques

Hi everyone,

Like Jiaxin and Johanna, I am also the ‘supervisor’ of the DRP students. Shahirah and me are taking the group that will be doing a project that involves proteases secreted by Stenotrophomonas maltophilia at 37^oC. We would have to teach them how to prepare the necessary media, teach them the things that they do not know (eg. how to streak in the proper way) and guide them through the experiment so that it is successful.

This is a very simple project and the purpose is to find out which are the strains that produce a lot of proteases. Strains that produce a lot of proteases might be more virulent as the proteases might be involved in the degradation of the host proteins involved in the immunity eg. Immunoglobulins (Ig).

Principle:

After growing the colonies, the colonies will be transferred onto milk agar. If the strain produces proteases, the proteases will digest the milk proteins and a clearing that looks like a halo around the colony will be observed.

Procedure:

Day 1 - Streaking of environmental strains

1. Streak the strains onto LB agar (Plate is divided into 2 as there are only 2 strains and only abit of the colony is needed)
   
   
2. Incubate the plate at 37^oC for 48 hours (As environmental strains do not grow very well at 37^oC, more time is required for them to grow)

Day 2 - Streaking of clinical strains, negative and positive control

1. Streak Escherichia coli (negative) and Pseudomonas fluorescens (positive) onto a LB agar each
   
   
2. Streak the strains onto LB agar (Plate is divided into 4)
   
   
3. Incubate at 37^oC for 24 hours

Day 3 - Transferring of colonies to milk agar

1. Divide and label the milk agar appropriately
   
   
   
2. Use a sterilised toothpick to touch the colony and then dab on the milk agar gently
   
   
3. Incubate the plates at 37^oC
   
   
4. Observe for halos (This will take about 3 days of incubation)

That’s all! Thanks=]

Chua Ming Boon

Tg01

0503197F

Florescence in situ hybridization

Florescence in situ hybridization (FISH), is a technique involving the separation of double stranded DNA by heat and hybridization of the target DNA sequence to the respective probes with a complementary sequence deirectly-labeled with a florochrome. The metaphases to which the probes anneal to are visualized as florescent signals of specific wavelengths under fluorescence microscopy. The number of signals observed in the metaphase spreads/interphase cells would indicate the copy number of the gene/gene loci concerned. This technique allows the identification of microdeleted/duplicated genes, aneuploidy screening (detection of extra chromosomes of chromosomes 13, 18, 21 X and Y), translocated genes in haematological malignancies and gene amplification from cases with cancer of the breast.

I would be explaining on identifying deleted/duplicated genes and loci that are related with microdeletion syndromes in congential disorders such as Prader-Willi/Angelman, Williams and Di-George using FISH techniques.

Samples – cultures of PHA(mitogen)-stimulated lymphocytes cells, amniotic fluids and chorionic villi.

Slide Preparation

1. Etch out the area on the underside of the glass slide with a diamond pen that contains the highest number of metaphase spreads. Slides can be kept in an airtight container at -20°C or 4°C if they are not used immediately.

2. Dehydrate the slide through an ethanol series (70%, 85% and 100% respectively) for 1 minutes each at room temperature. Blow-dry the slide with a hair dryer.

Probe Preparation and hybridization
The following steps must be carried out in reduced lighting as the probes are light-sensitive.

1. Remove the probe vial from the freezer and thaw at room temperature. Centrifuge the vial.

2. Add probe mixture onto the etched area of the slide and cover with a circular glass coverslip.

3. Seal the edges with rubber cement and place the slide into a light proof box.

4. Switch on the 'HYBrite' system and set the desired program (denaturation and hybridization steps).

5. Moisten the toweling paper in the troughs with water. Place the slide onto the hybridization surface, coverslip facing upwards.

6. Ensure that the target area is in contact with the hybridization surfaces and also that all the remaining slide positions are filled with blank slides to even out the temperature.

7. Close the 'HYBrite' lid and commence the program.

Post Hybridization Wash
Carry out the following steps in reduced lighting.

1. Switch on water bath. Set the temperature to 75° C. Prepare two coplin jars for Wash A and Wash B or two plastic slide chambers if only 1 or 2 slides are required for washing.

2. Place the coplin jar/slide chamber containing Wash A in to the water bath when it is just switched on.

3. Leave the coplin jar/slide chamber containing Wash B at room temperature.

4. When the water bath reaches its set temperature, remove the slide from the 'HYBrite' system. Peel off the rubber cement from the coverslip with a pair of forceps. Remove the coverslip.

5. Place the slide in the Wash A and agitate the slide by jiggling briefly for about 3 seconds. Wash for exactly 2 minutes.

6. Remove the slide and transfer to Wash B. Agitate the slide by jiggling briefly for about 3 minutes. Wash for 1 minute.

7. Remove the slide and air or blow dry completely in the dark at room temperature. Apply 10m l of DAPI counterstain (Vectashield mounting medium) and mount a glass coverslip over the hybridize area. Seal with nail vanish.

Image of Digeorge Syndrome

Adapted from :
http://www.pathology.washington.edu/galleries/Cytogallery/main.php?file=intro

Image of HYBrite system

Adapted from:

http://www.ebiotrade.com/buyf/productsf/vysis/img189_1.jpg

Reference:

American College of Medical Genetics Laboratory (1999). Standard guidelines for clinical genetics laboratories, 2nd Ed. Bethesda, MD: ACMG.

By:

Yvonne Lau

0503149G

TG01

Lab Techniques

HELLO! =D

If you're wondering what we the in-house students are doing for SIP. NO NO.. not preparing reagents and stock checks! For our group, our assigned task is to be the "supervisors" of the DRP (Differential Research Programs) students!

Johanna and I will be taking one of two groups who are under Dr. Quek's project. This project involves extraction of periplasmic proteins using chloroform (CHCl₃) shock, of an isolate of Stenotrophomonas malitophilia which is incubated at two different temperature 28°C (environmental) and 37°C(human body temp.). The differences in protein expression will be analyzed and compared. Experiment will be repeated on other isolates too. This is also the continuation of what Jo and I have been doing for our last DRP.

Outline of Workflow/Protocol

Day 1 (Streaking)

1. Streak the different isolates on a well divided LB Agar plate
2. Incuabte plates for 24 hours at 37°C

Day 2 (Inoculation)

1. All colonies of each isolates is scrapped off and mix in 5mL PBS
2. Cell density (Opitcal Density 600) taken in triplicates to determine number of cell present
3. Volume of inoculum equivalent to 5.0X107 is inoculated into 2 tubes
4. One tube incubate at 28°C and the other at 37°C for 16 hours 250 rpm

Day 3 (Extraction)

1. After 16 hours, the tubes are spin down at 3000xg for 20 minutes at 10°C

2. Decant the supernatant and resuspend the cells in 10mL of PBS

3. Spin down the tubes at 3000xg for 20 minutes at 10°C

4. Repeat steps 2 and 3.

5. Decant the supernatant and resuspend the cells in 10mL of PBS

6. Take OD600 reading in triplicates with 10X dilution

7. Calculate the volume withdrawal equivalent to 1010 number of cells

8. Pipette the calculated volume into 1 microfuge tube

9. Spin down cells at 3000xg for 20 minutes at 10°C

10. Decant supernatant

11. Resuspend pellet in remaining medium by brief vortexing

12. Add 20mL of chloroform to each tube

13. Incubate at room temperature for 15 mintues

14. Add 200uL of 0.1M Tris-HCl to tubes to stop reaction

15. Spin tubes at 6000xg for 20 minutes at 10°C

16. Collect the supernatant to a new microfuge tube

17. Spin the supernatant tubes at 14000xg for 10 minutes at 10°C

- This is to ensure that only the periplasmic proteins are present in the supernatant as cell debris will be pellet down

18. The supernatant is pipetted out into a new microfuge tube

Example of how to calculate 5.0X107 worth of cells

Average OD600 Reading = 1.70

Desired number of cells = 5.0X107

1.00 OD = 1.4X107 cfu/mL

Volume of inoculum = (Desired no. of cell) / (Average OD600 X 1.4X107) X1000

= (5.0X107) / (1.70 X 1.4x107)

= 21.0uL

After extraction, protein concentration will be determined by Bradford assay. The samples will be run on 1-D gel (separation by MW) and stained with either Coomassive Blue or Silver stain. Protein bands will be analyse and compare to identify the difference in protein expression of an isolate incubated at 28°C and 37°C.

Post Stained 1D Gel
Adapted from: http://www.raytest.de/bio_imaging/applications/gel_documentation/applications_geldoc1.jpg

As this is a project that lasts for only 3 weeks, the job load is lesser and simpler.

That's all. Have great day!

Tang Jiaxin TGo1

0503257H

Lab Technique

Hiya...I shall talk about purification of primers and followed by DNA sequencing after finding the optimization temperature,a lab technique that I have done after PCR. The purpose of purification is to ensure that the unwanted unbound stuff like unbound primers or excess buffer will not be present and thus will not affect the results after DNA sequencing is done. Feel free to ask any questions ya..

Purification of PCR Products:

Materials

Shrimp Alkaline phosphotase (SAP)

Exonuclease 1 (Exo 1)

Method:

1) Prepare a master-mix containing both the enzymes for purification. (Refer to table below for purification reaction mixture set-up.)

Purification reaction mixture set-up

Components Required:	Volume Required/ul:	Master-mix Required/ul:
SAP	1.0	80
Exo 1	0.5	40
	1.5	120

2) Pipette 15ul into 8 eppendof tubes. Using a multi-channel, pipette 1.5ul of the master-mix into the wells of the PCR plates.

3) Place the PCR plates into the machine.

DNA Sequencing (Cycle sequencing):

Materials:

Sequencing dye

Sequencing buffer

Forward primer

Reverse primer

Autoclaved water

Method:

1) Prepare a master-mix containing all the components for cycle sequencing. (Refer to table below for cycle sequencing reaction mixture set up.)

Cycle sequencing reaction mixture set up

Components required:	Volume Required/ul:	Master-mix Required/ul:
Sequencing dye	1.0	80.0
Sequencing buffer	0.5	40.0
Forward primer OR reverse primer	0.4	32.0
Autoclaved water	0.6	48.0
	2.5	200

2) Pipette 2.5ul of the required reaction mixture into a new PCR plate.

3) Pipette 2.5ul of the correct purified PCR products into the new PCR plate, which contains the reaction mixture. Ensure it is mix well.

4) Spin down the new PCR plates.

5) Place the new PCR plates into the machine.

DNA Sequencing (After cycle sequencing):

Materials:

3M of Sodium acetate

125mM of EDTA

100% Ethanol

70% Ethanol

Formamide

Method:

1) Spin down the PCR products in the PCR plates.

2) Prepare a master-mix of the stock solutions (200ul of 3M sodium acetate and 200ul of 125mM of EDTA).

3) Pipette 2ul of master-mix of the stock solutions into each well of the PCR plate, using a multi-channel.

4) Dispense 25ul of 100% ethanol into each well of the PCR plat, using a multi-channel.

5) Pipette everything from the PCR plates and transfer them into sequencing plates, using a multi-channel.

6) Wrap the sequencing plates with aluminum foil and incubate them at room temperature for 15 minutes. Dispose the PCR plates.

7) Centrifuge the sequencing plates at 3000G at 4^oC for 25 minutes.

8) Invert the sequencing plates using 3 pieces of tissue paper and pulse them at 200G at 4⁰C for 20 seconds.

9) Dispense 70% ethanol into each well of the sequencing plates.

10) Centrifuge the sequencing plates at 1700G at 4^oC for 15 minutes.

11) Invert the sequencing plates using 4 pieces of tissue paper and pulse them at 200G at 4^oC for 20 seconds.

12) Repeat step 11, using the other sides of the tissue paper.

13) Place the sequencing plates into a vacuum dryer, not more than 7 minutes.

14) Pipette 20ul of formamide into each well of the sequencing plate/

15) Cover the sequencing plates with rubber mats and place the sequencing plates in the septa. Place them into the sequencer machine.

Michelle TG02 0503808H

Lab Techniques

CyDye Fluor in DIGE

Hi guys! Hope all of you are doing well in the 10th week of your attachment, 10 more weeks to go so hang in there all right. For those who are enjoying your work, appreciate every week cause all good things must come to an end :p

For this blog entry, I will be sharing with you guys about CyDye used in DIGE. DIGE stands for DIfferential Gel Electrophoresis. It is a technique used to detect and identify proteins involving 2D-gel electrophoresis (scroll down to Jiaxin’s entry for more info). It allows up to three different protein samples to be separated on the same gel. This ability is termed multiplexing. Each protein sample is labeled with one type of dye i.e. CyDye in this case.

Now what is CyDye then? Basically it is a cyanine, fluorescent dye used to label the proteins. There are different forms of CyDye depending on the sample you are dealing with. For example there is a set of CyDye to label scarce samples. Since the sample is limited, a lot of dye is added and bind to the proteins at certain amino acid residues such that they become saturated with the dye. The saturation will help in the detection of the presence of the proteins after 2D- gel electrophoresis. In general, CyDye is excited lasers at specific wavelengths and emits a signal of a narrow wavelength range. There are three CyDyes used in most experiments; Cy2, Cy3 and Cy5. Cy2 is excited by a laser at a wavelength of 488nm whereas Cy3 is excited at 532nm and Cy5 at 635nm.

For my group’s experiments, we will most probably use one of the three dyes i.e. Cy3. We are only using one as we intending to test if this method of labeling is suitable for the bacterium that we are working on i.e. Stenotrophomonas maltophilia. At the moment we are trying out a protocol to extract only the exposed cell surface proteins of this gram-negative bacterium and we are using this dye to label the surface membrane proteins.

As mentioned earlier that there are various forms of cyanine dyes depending on the experiment. The Cy3 that we are using is a minimal dye. This dye has an ester group on it. It will bind only to the epsilon amino group of lysine found on proteins to form an amide linkage. Most proteins have a number of lysine residues on them thus these proteins will be labeled with the cyanine dye. It is termed a minimal dye as it is limiting in the reaction between the proteins and the dye. Only a small amount of the dye will be added to ensure that only 1-2 % of the available lysine is labeled with the dye. This is so as during identification of the proteins using mass spectrometry like MALDI (refer to my previous post), the proteins will be cut using trypsin. Trypsin cleaves at lysine. Thus, we use minimal labeling such that it will not affect the mass spectrometry results. Once the proteins are labeled, the sample will be subjected to gel electrophoresis. For my experiment, the sample will be run on 1D-gel where the separation of proteins is based solely on its molecular weight.

That will be all for my post. Take care and say hi to week 11!

Posted by: Shahirah Bibi
0503174E
TG01