C-banding of Fish Chromosomes
Because high-resolution banding is difficult for fish chromosomes, comparative information mostly drawn from fish karyotyping is the number of chromosomes and the arm number. We feel frequently, however, difficulty to distinguish mono-armed chromosomes from bi-armed ones. Which should we classify sub-telocentric chromosomes as mono-armed or bi-armed?. C-banding technique could answer this question.
C-banding technique is rarely applied in recent banded karyotype studies of fishes, while many studies employ Ag-NOR and CMA3 staining techniques to visualize nucleolar organizing regions. Scientists might misunderstand that C-banding is a method for simple localization of centromeres which is easily identified by the conventional staining and that laborious C-banding is no more profitable to them. C-banding could visualize, however, not only centromeres but also distinction between euchromatic and heterochromatic short arms of sub-meta to sub-telocentric chromosomes. If the entire short arm of a chromosome is heterochromatic, that chromosome is a modified acrocentric element with a redundant heterochromatic block. We could thus determine the precise number of chromosome arms by C-banding. Here I present methods to visualize C-heterochromatin in fish.
Three pairs of Giemsa stained fish chromosomes. Left and center, chromosomes possibly identified as sub-telocentric. Right, a 'metacentric' chromosome pair.
Sequential C-banding of the same chromosomes revealed their real configuration. Left, a sub-telocentric pair with a dot-like cetromeric heterochromatic block and a euchromatic short arm. This chromosome pair should be classified as bi-armed. Center and right, chromosome pairs with heterochromatic short arms. These chromosomes are acrocentric with redundant heterochromatin and are hereby regarded as mono-armed chromosomes.
ROUTINE GIEMSA STAINING OF CHROMOSOME SPREADS
The method I present here is a sequential staining technique of the routine Giemsa staining and C-banding. Because C-banding makes chromosomes to swell, routine Giemsa staining prior to C-banding is useful to observe gross morphology of chromosomes.
Chromosome spreads are made by routine air drying method. Flame drying is not suitable. Adjust the condition of the cells in which chromosomes weakly condense with sister chromatids stick to each other (see above). Try a few patterns of colchicinization and hypotonization to find out the best condition for your species and samples.
Stain the slides for 10min in 2% Giemsa (Merck) in 1/15M Soerensen buffer (pH5.9). Wash briefly with distilled water and jet dry. Do not apply cover glass.
Take photos of chromosome spreads of good condition with an oil-mediated 100x objective lens of a microscope. Record the position of the spreads with the micrometer on the stage of the microscope. If you use a classical film and photo-paper system, use the Minicopy HRII film (Fuji) which is developed with the Copinal (Fuji) and print on #4 contrast photo-papers. Take photos with film sensitivity set at ISO32 and 2/3x - 1/2x underexposure. For printing, dodge and burning around inner and outer portion of cell spreads are advisable.
DESTAINING
Freshness (within about 7days after air-dry) of slides is important.
Soak slides for 20min in xylene to remove the oil. Complete removal of the oil is important. Fresh xylene in a larger staining jar (c.a. 100ml) can degrease about 10 slides. Change of fresh xylene may give better results. Jet dry. You can check if the oil is completely removed by gentle breezing onto the slide. If the slide is uniformly steamed, it is OK.
Soak slides for 10-20min in 70% ethanol to remove Giemsa. Dye cannot be removed completely, but it is OK. Jet dry. You can also check here if the oil is completely removed. If the slide is repellent to alcohol, dip it again in xylene to ensure complete degrease.
ALKALI DENATURE AND ANNEALING OF CHROMOSOMAL DNA
Soak slides for 50-100sec in saturated Ba(OH)2 solution at 50oC, and then dip them in 0.1N HCl at room temperature.
Duration of the alkali treatment depends on the condition of chromosome spreads and also different from species to species. For mitotic spreads of cobitine loach, it would be 90-100sec, while 80-90sec treatment is good for meiotic spreads of these species. Try a few patterns of alkali treatment to find out the best condition for your species and samples. If the entire chromosome is stained uniformly dark, prolong this step. On the other hand, if chromosome swells too much and is stained uniformly light, shorter duration may be good.
When slides are put in the HCl solution, powdery alkalic matter stuck onto the slides rapidly dissolves. Within a minutes, take out the slides, wash briefly with distilled water, jet dry, and incubate them in 2xSSC at 60oC for 1hr.
RESTAINING
Take out the slides from 2xSSC, wash briefly with distilled water, and jet dry. Stain the slides with 4% Giemsa in 1/15M Soerensen buffer (pH6.8) for 20min. Wash briefly with distilled water and jet dry. Mount the slides again onto the microscope stage, dial to the positions you recorded, and take photos of the C-banded chromosome spreads. For use of a classical film and photo-paper system, the Minicopy HRII film and #3 contrast photo-papers are suitable. Take photos with film sensitivity set at ISO32 and 1x - 2/3x exposure time.
AN ALTERNATIVE METHOD
If the chromosomes are too much elongated and only lightly stained with the routine Giemsa staining, soak the slides in 0.1N HCl for 20min at room temperature prior to the alkali treatment. In this case the alkali treatment is shorten to 45-50sec (for cobitine loach). After the alkali treatment, dip the slides again briefly in the HCl solution and then in 2xSSC as usual.
REFERENCE
Saitoh K. 1986. A preliminary note on chromosomes of F 1 hybrid between middle and small races of the striated spined loach (Cobitis taenia striata). Ann Rep Biwako Bunkakan 4:62-65.
Saitoh K. 2003. Mitotic and meiotic analyses of the 'large race' of Cobitis striata, a polyploid spined loach of hybrid origin. Folia Biol (Krakow) 51(Suppl):101-105.
CHEMICALS AND SOLUTIONS
Soerensen Buffer (pH5.9)
For 1000mL
Na2HPO4.2H2O 1.19g or Na2HPO4.12H2O 2.32g
KH2PO4 8.19g
Add distilled water up to 1000mL. Stable for one year under refrigeration.
Soerensen Buffer (pH6.8)
For 1000mL
Na2HPO4.2H2O 5.95g or Na2HPO4.12H2O 11.6g
KH2PO4 4.55g
Add distilled water up to 1000mL. Stable for one year under refrigeration.
To prepare 2(4)% Giemsa solution, add 0.6(1.2)mL Giemsa stock solution (Merck) to 30-35mL Soerensen buffer. Prepare the staining solution freshly everyday. Giemsa dye is hydrophobic and undissolved dye floats on the buffer surface. This makes dye blobs on the slide glass. To avoid this, put a cut paper towel on the buffer surface to stick out the floating dye, or mix vigorously making bubbles with a pipette.
Saturated Ba(OH)2 Solution
Freshly prepare the alkali solution every time. To do this, pre-warm water in a small staining jar (c.a. 35ml) to 55oC in a water bath. Take out the jar from the bath. Put a spoonful of crystal Ba(OH)2 into the warm water, mix briefly, and make sure the water temperature drops to about 52-53oC. Putting a few glass slides at room temperature makes the water temperature down to 50oC. Store crystal Ba(OH)2 tightly capped (avoid exposure to CO2 in the air).
0.1N HCl
For 1000mL
Conc HCl 9.8mL
Add distilled water up to 1000mL. Stable for more than one year at room temperature.
2xSSC
Make 20xSSC as stock solution and dilute 10 times freshly everytime for use.
20xSSC for 1000mL
NaCL 175.32g (3M)
Na3-citrate.2H2O 88.23g (0.3M)
Add milli-Q water up to 1000mL. Stable for more than one year at room temperature.
