• A series of rapid nuclear cycles takes place during the first few hours of Drosophila development, some of cycles take as little as 10 minutes. This high mitotic activity provides the early Drosophila embryo a very useful system to study various aspects of cell cycle and chromosome behaviour.
• For the study of these processes, it is essential to apply immunofluorescence-based techniques to visualize the subcellular organelles or the molecules under study.
• Apart from its own cell membrane, the Drosophila embryo is enclosed by the vitelline membrane and the chorion.
• These membranes being impermeable to most chemicals, it must be removed for the fixatives to penetrate the embryo.
• The removal of the chorion is technically very simple and is harmless to the embryo.
• The removal of the vitelline membrane can be done only after fixation of the embryo.
• The need to sort out this understanding between vitelline membrane removal and embryo fixation is the character specific to the protocols for immunostaining Drosophila embryos.
• Even if the basic procedure remains unchanged, many of the details of the protocols for immunostaining embryos have been modified, and others have been removed altogether.

Protocol for immunostaining Drosophila embryos.

1. Allow flies to lay eggs on 3 % agar plates containing 4 % fruit juice.
2. Brush the embryos from the collecting trays in 0.7 % saline and place them on a nylon gauze in a Millipore filtration funnel.
3. Remove the chorion by passing commercial bleach over the embryos for 3 minutes.
4. Wash thoroughly with water.
   • The embryos can be processed immediately after any of the fixation protocols described later OR, they can be poured onto a dry plastic Petri dish, and covered with water so that they can be kept alive and at the same time be easily observed under the dissection micro- scope. This allows the selection of those embryos that have reached a particular developmental stage.
5. If formaldehyde fixation (4%or 37%) is chosen, proceed to step 5. If methanol fixative is to be used, go directly to step 6.
6. Transfer the embryos into a glass vial containing 1 ml of either 4 % or 37 % formaldehyde and 4 ml n-heptane. Incubate in spinning wheel for 20 minutes.
7. Place the embryos into a microcentrifuge tube containing 500 methanol and 500µl heptane. Invert the tube several times. Most embryos will lose their vitelline membrane and sink to the bottom of the vial.
8. Remove all heptane and as much methanol as possible.
9. Add fresh methanol.
   • If the embryos were not fixed with formaldehyde, keep them in methanol for a further 2 hours at room temperature or overnight at 4°C.
10.  Rehydrate in phosphate-buffered saline (PBS)
11. Block any residual fixative by incubating the embryos in 10% fetal calf serum (FCS), 0.3% Tween in PBS for 1 hour.
    • If RNase treatment is to be carried out, an aliquot of a boiled solution of the enzyme should be added at this time at a concentration of 2 mg/ml.
12. Incubate the preparation with the primary antibody in 10% FCS, 0.1% Tween in PBS either at 4°C overnight, or for 4 hours at room temperature.
13. Wash several times with 0.1% tween in PBS over a I-hour period.
14. Incubate with the secondary antibody for 4 hours at room temperature or overnight at 4°C.
15. Wash as before, but using PBS alone for the last two or three washes.
16. To mount, place a drop of approximately 50 µl of mounting medium on a microscope slide and transfer the embryos into the drop. Move them around with a pair of twizers until they get embedded, cover with a coverslip, and seal the preparation with nail varnish.

Immunostaining of developing Embryos of Drosophila

The post Immunostaining of developing Embryos of Drosophila appeared first on Online Biology Notes.

• The larval brain is the tissue of choice for the study of cell division during late development. Most mitotic mutants of Drosophila have been identified by screening squashed preparations of third-instar larval brains.
• We include here two protocols for immunofluorescent staining cells from the larval brain.

lmmunfluorescent staining by Squashed Preparations of larval Brains:

1. Dissect out larval brains in 0.7% NaCl.
2. Place the brain in a microcentrifuge tube containing 3.7% formaldehyde in phosphate-buffered saline (PBS), and incubate at room temperature for 1 hour.
3. Remove the formaldehyde, add 10% Fetal calf serum (FCS), 0.3%triton X-100 in PBS, and incubate for 1 hour at room temperature.
4. Place the brains on a microscope slide, add a drop of 45% acetic acid, and leave for 30 seconds.
5. Remove the liquid using tissue paper, add a drop of 60% acetic acid, and cover with an 18 X 18 mm2 coverslip.
   • Do not squash yet, but wait for 3 minutes.
6. Squash between two sheets of blotting paper by pressing with two fingers on opposite comers.
   • Keep pressing for at least 10 seconds and release pressure gently.
   • Repeat, pressing on the other two comers.
7. Immerse the end of the slide carrying the coverslip in liquid N₂.
   • When the nitrogen ceases to boil, remove the slide and lever off the coverslip with the flick of a scalpel.
8. Dehydrate by successive immersion of the slides in 70% ethanol for 3 minutes, 100% ethanol for 3 minutes, and air dry before use.
9. Carry out immunostaining as described for in toto preparations.

lmmunostaining Whole-Mount Preparations of larval Brains:

1. Dissect out larval brains in 0.7% NaCl.
2. Incubate for 30 minutes in 3.7% formaldehyde followed by 60 minutes in 37% formaldehyde.
3. Transfer to 0.3% triton X-100, 10% FCS, in 0.7% NaCl, and keep there for about 20 minutes.
4. Incubate with the first antibody in 2.5 mg/ml RNAase, 0.3% triton X-100, 10% FCS, in 0.7% NaCl overnight
5. Wash in 0.3% triton X-100, 10% FCS, in 0.7% NaCl at least four times for 10 minutes.
6. Incubate with second antibody in 0.3% triton X-100, 10% FCS, in 0.7% NaCl at least for 2 hours.
7. Wash in 0.3% triton X-100, 10% FCS, in 0.7% NaCl at least four times for 10 minutes, and then in 10% FCS in 0.7% NaCl for a few minutes.
8. If required, incubate in propidium iodide 1in 0.7% NaCl for 5 minutes.
9. Transfer the brains into a drop of approximately 50 pl of mounting medium on a microscope slide, move them around with a pair of twizers until they are embedded, cover, and seal with nail varnish.

Immunofluorescent staining of Drosophila Larval Neuroblasts

The post Immunofluorescent staining of Drosophila Larval Neuroblasts appeared first on Online Biology Notes.

• In situ hybridization to diploid cells serves in mapping and characterizing the behaviour of heterochromatic sequences of Drosophila, which are both under represented and aggregated in polytene cells.
•  Two protocols are included for in situ hybridization to diploid cells from squashed and whole-mounted third-instar larval brains.
  • For genome mapping, squashed preparations are preferred as they are easy to make and provide the highest possible resolution.
  • The procedure can be employed either in untreated brains or in brains where the number of mitotic chromosomes has been artificially increased by incubation with colchicine.
  • Whole mounted protocol should be carried out when more functional studies are to be performed .
  • Whole-mounted tissues conserve the original three-dimensional arrangement of subcellular organelles and are devoid of the artefacts produced by squashing.
  • These are necessary requirements needed to study problems like chromosome pairing, chromosomal domains in interphase nuclei, attachment sites, and many others.

Fluorescent in-situ Hybridization to Squashed Diploid Cells:

1. Obtaining brain of larvae:
   • With a pair of tweezers, brains are obtained from third-instar larvae by pulling from the mouth parts while holding the larvae by their middle.
   • The brain (ventral ganglion plus optic lobes) usually comes out together with the salivary glands and other internal tissues.
   • Carefully remove other tissues as brain preparations devoid of other tissues are essential to obtain good squashes.
2. Dissect out larval brains in 0.7% NaCl and incubate in 0.5 kg/ml colchicine in 0.7% NaCl in a dark, humid chamber for 2 hours.
3. Apply hypotonic shock by washing in 0.5% trisodium citrate for 10 minutes.
4. Place the dissected brains on a microscope slide, add a drop of 45% acetic acid, and leave for 30 seconds.
5. Remove the liquid using tissue paper, add a drop of 60% acetic acid, and cover with a 18 X 18 mm² coverslip.
   • Do not squash yet, but wait for 3 minutes.
6. Squash between two sheets of blotting paper by pressing with two fingers on opposite comers.
7. Keep pressing for at least 10 seconds and release pressure gently. Repeat, pressing on the other two comers.
8. Immerse the end of the slide carrying the coverslip in liquid N₂.
   • When the nitrogen ceases to boil, remove the slide and level off the coverslip with the flick of a scalpel.
9. Dehydrate by successive immersion of the slide in 70% ethanol for 3 minutes, 100 %ethanol for 3 minutes, and air dry before use.
10. Bake the slide at 58°C for 1 hour in a dry oven.
11. Immerse slides in H₂O gently to remove the coverslips.
12.  Denature the chromosomal DNA by boiling for 3 minutes.
    • The slides should be kept in hot (>80°C) water until the probe is applied.
13. Apply 10 µl denatured probe and carry out the hybridization at 58°C in a humid chamber overnight
14.  After hybridization, carry out the washing and Fluorescein isothiocyanate (FITC) staining.

Fluorescent in Situ Hybridization to Whole-Mounted Diploid Tissues

1. Dissect brains in saline.
2. Fix for 10 minutes in formaldehyde 3.7% in saline.
3. Transfer to an Eppendorf tube and add about 500µl of 37% formaldehyde and 200 µl of n-heptane.
4. Incubate for 20 minutes in spinning wheel at room temperature.
5. Remove all the heptane and as much formaldehyde as possible, add 500 µl of 1% triton X-100 in PBS, and incubate for 20 minutes.
6. Remove supernatant and wash once in phosphate-buffered saline (PBS)
7. Add 50 of probe and cover with 50 parafilm oil.
8. Denature in boiling water bath for 10 minutes and incubate overnight at 58°C.
9. Add 4X saline sodium citrate (SSC)  under the oil and remove as much oil as possible.
10. Wash for 10 minutes in 4X SSC.
11. Wash for 10 minutes 4X SSC + 0.1% triton X-100.
12. Wash for 10 minutes in 4X SSC.
13. Incubate in 150 FITC-avidin in 4X SSC for 30 minutes at room temperature and repeat washes as in steps 10 to 12.
14. Incubate in 1 propidium iodide in 4 X SSC for 10 minutes.
15. Wash for 10 minutes in 4X SSC.
16. Mount in glycerol-propyl gallate.

Fluorescent in-situ Hybridization to Diploid Cells

The post Fluorescent in-situ Hybridization to Diploid Cells appeared first on Online Biology Notes.

1. Dissect out salivary glands from third-instar larvae in 0.7% NaCl.
2. Transfer the salivary glands to a drop of 45 % acetic acid for 30 seconds and then transfer to a 1:2:3 mixture of lactic acid: glacial acetic acid: water and incubate for 5 minutes.
3. Cover with a siliconized coverslip.
4.  Sandwich between two sheets of blotting paper, and tap with the blunt end of a pair of tweezers to squash the cells.
5. Observe the squashing procedure by analyzing the chromosomes by phase-contrast microscopy (a 40X dry objective is ideal for this purpose).
6. When the chromosome spreading is optimum, leave to fix for l to 2 hours at room temperature or overnight at 4°C.
7. To remove the coverslip, immerse the preparation in liquid N, until boiling stops, level off the coverslip with the flick of a scalpel.
8. Immerse the slide immediately in a jar containing 70% ethanol for at least 3 minutes.
9. Slides can be kept at this stage for a long time. This is an appropriate step at which to accumulate all the slides so that they all can go into the next steps at the same time.
10. When all the slides are ready, they are transferred into absolute ethanol for 3 minutes and air dried.
11. Before denaturation, the slides are treated by immersion in a jar containing 2X saline sodium citrate (SSC) at 65°C for 30 minutes, followed by two immersions in 70% ethanol for 5 minutes and absolute ethanol for 5 minutes and air dried as before.
12. Place the slides in freshly made 70 mM NaOH for 3 minutes in order to denature DNA.
    • It is very important that this solution is made fresh every time.
13. Transfer the slides to 70% ethanol for 3 minutes, absolute ethanol for 3 minutes, and air dry as before.
14. Place 20 of the denatured probe on top of the chromosomes and cover with a coverslip.
15.  Hybridization is carried out in a humid chamber overnight at 58°C.
16. After hybridization the slides are washed in the following series:
    • 2X SSC at 53°Cfor 2 minutes. Check that the coverslip falls off.
    • 4X SSC at room temperature for 5 minutes.
    • 4X SSC containing 0.1% triton X-100at room temperature for 5 minutes.
    • 4X SSC at room temperature for 5 minutes. Make sure that the last wash does not contain any triton X-100. These washing solutions can be kept and reused.
17. Incubate the slides in 2% ( v/v ) fluorescein isothiocyanate (FITC)- labelled avidin in phosphate-buffered saline (PBS) for 30 minutes.
    • Wash in the following series:
    • 4X SSC at room temperature for 5 minutes.
    • 4 X SSC containing 0.1% triton X-100 at room temperature for 5 minutes.
    • 4X SSC at room temperature for 5 minutes. Make sure that the last wash does not contain any triton X-100.
    • If necessary, the signal can be enhanced 5- to 10-fold by incubation for 30 minutes with 1:100 FITC anti-avidin D followed by the same series of washes.
18. The preparation is now ready to be mounted for microscope examination.
19.  Remove the excess liquid with a paper tissue and add a drop of mounting medium containing 1 g/ml propidium iodide to counterstain DNA.
    • There are several mounting media that can be used.
    •  The simplest one is a solution of 2.5% propyl gallate in 85% glycerol.
    • This solution is economical and easy to make, but it does not set, thus needs the coverslip to be sealed.
    • This can be gained by applying nail varnish on the edges.
    • Other mounting mediums such as Permount or Gelvatol can also be used.

Preparation of Polytene Chromosomes of Drosophila for In-Situ Hybridization

The post Preparation of Polytene Chromosomes of Drosophila for In-Situ Hybridization appeared first on Online Biology Notes.

• The modification is seen in the detection system, which is based on a fluorescence signal instead of the coloured precipitate resulted by an enzymatic reaction.
• Advantages: Fluorescent system has two major advantages that make up for this relatively minor difficulty.
  • The first advantage is a observable increase in sensitivity. It is possible to produce preparations in which the background fluorescence signal is virtually null under optimal hybridization conditions, thus achieving high signal-to-noise ratios.
  • The combination of such high- quality preparations with any of the highly sophisticated fluorescence microscope systems, such as confocal microscopes or cool charge-coupled device (CCD)cameras, can detect accurately in comparison to enzymatic detection methods.
  • The second advantage is the enhanced resolution. This is specially useful when more than one target sequence is being analysed at the same time.
  • The fluorescent protocol can be employed for double or triple hybridization with multiple probes that can easily be detected using the appropriate non-overlapping fluorochromes.
  • In such case, the location of the different target sequences can be determined without the constraints inherent to transmitted light microscopy.
• limitations:
  • The main limitation of this protocol is the short-lived nature of the fluorescent signal.
  • Unlike the products of enzymatic reactions, fluorescence vanishes away over time, and bleaches out rapidly when observed under the microscope.
  • Therefore, fluorescence-based preparations are temporary.

Preparation of the Probe for In-situ Hybridization

• Mix up the reaction mixture containing:
  • 5µl of 5 X oligo-labelling buffer minus dT (deoxy thymine)
  • 1 µl of biotin-16-uridine triphosphate approximately 100 ng of DNA (boiled)
  • 1 Klenow fragment of DNA polymerase (1 unit)
• Make up to a final volume of 25µl with water.
• Leave it for about 2 hours at 37°C or overnight at room temperature for effficient labelling
  • There is no need to remove unincorporated nucleotides because of the very high incorporation rates.
• Store this probe solution at -20°C for many months.
• Before hybridization, the probe should be diluted in 1 volume of water and 2 volumes of 2X hybridization buffer.
• Then it should be boiled for 3 minutes and quenched on ice before it is applied to the chromosomes.

Fluorescent in-situ hybridization: advantages, limitations and preparation of probe

The post Fluorescent in-situ hybridization: advantages, limitations and preparation of probe appeared first on Online Biology Notes.

Materials required:

• Overnight culture of Staphylococcus aureus in LB
• TE buffer ( 10mM tris.Cl, 1mM EDTA, pH 8.0)
• 10% SDS
• 1% lysozyme
• 1:1 Phenol-Chloroform mixture
• Chloroform
• 5N NaCl
• 5M Ammoniun acetate
• Ice cold isopropanol
• 70% ethanol

Protocol of DNA extraction from gram Positive bacteria:

1. Take 1.5 ml of bacterial broth culture (overnight culture in LB) into a microfuge tube.
2. Centrifuge at 800rpm for 10 minutes at 4°C and discard the supernatant.
3. Suspend the pellet in 400µl TE buffer. Mix well by vortexing.
4. Add 20µl of 1% lysozyme, mix well and incubate at 37° for 10 minutes in water bath.
5. Add 30µl of 10% SDS and mix it.
6. Incubate the tube at 37°C for 30 minutes in water bath.
7. Shear the cell suspension 3-5 times with the help of 26G needle.
8. Add 500µlof 1:1 phenol-chloroform mixture.
9. Centrifuge at 13000rpm for 2 minutes at 4°
10. Transfer the supernatant into another microfuge tube.
11. Add 500µl of chloroform and centrifuge at 13000rpm for 2 minutes at 4°
12. Transfer the supernatant into another microfuge tube.
13. Add 25µl of 5N NaCl and centrifuge at 13000rpm for 10 minutes at 4°
14. Discard the supernatant and suspend the pellet in 100µl TE buffer.
15. Incubate the tube at 37°C for 30 minutes.
16. Add 50 µl of 5M ammonium acetate.
17. Add 250 µl of cold isopropanol and incubate at room temperature for 5 minutes.
18. Centrifuge at 13000rpm for 5 minutes at 4°
19. Discard the supernatant and wash the pellet with 100 µl of 70% ethanol.
20. Pour off the ethanol and invert the tube on a clean absorbent paper to drain.
21. Allow the pellet to air dry for 5-10 minutes.
22. Suspend the pellet in 100µl TE buffer.
23. Store at -20°

DNA extraction from Gram Positive bacteria (Staphylococcus aureus): materials required and protocol

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Materials required:

• Overnight culture of coli in LB
• TE buffer ( 10mM tris.Cl, 1mM EDTA, pH 8.0)
• 10% SDS
• 1:1 Phenol-Chloroform mixture
• Chloroform
• 5N NaCl
• 5M Ammoniun acetate
• Ice cold isopropanol
• 70% ethanol

Protocol of DNA extraction from E.coli:

1. Take 1.5 ml of bacterial broth culture (overnight culture of coli in LB) into a microfuge tube.
2. Centrifuge at 800rpm for 10 minutes at 4°C and discard the supernatant.
3. Suspend the pellet in 400µl TE buffer. Mix well by vortexing.
4. Add 10µl of 10% SDS and mix it.
5. Incubate the tube at 37°C for 1 hour in water bath.
6. Shear the cell suspension 3-5 times with the help of 26G needle.
7. Add 500 µlof 1:1 phenol-chloroform mixture.
8. Centrifuge at 13000rpm for 2 minutes at 4°
9. Transfer the supernatant into another microfuge tube.
10. Add 500 µl of chloroform and centrifuge at 13000rpm for 2 minutes at 4°
11. Transfer the supernatant into another microfuge tube.
12. Add 25µl of 5N NaCl and centrifuge at 13000rpm for 10 minutes at 4°
13. Discard the supernatant and suspend the pellet in 100µl TE buffer.
14. Incubate the tube at 37°C for 30 minutes.
15. Add 10 µl of 5M ammonium acetate.
16. Add 250 µl of cold isopropanol and incubate at room temperature for 5 minutes.
17. Centrifuge at 13000rpm for 10 minutes at 4°
18. Discard the supernatant and wash the pellet with 100 µl of 70% ethanol.
19. Pour off the ethanol and invert the tube on a clean absorbent paper to drain.
20. Allow the pellet to air dry for 5-10 minutes.
21. Suspend the pellet in 100µl TE buffer.
22. Store at -20°

DNA extraction from E. coli: materials required and protocol

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