The aim of clinical chromosome analysis is to determine whether or not a horse has a normal set of chromosomes. The analysis is usually carried out on animals with various reproductive and/or developmental disorders to ascertain possible causes of abnormalities. Additionally, cytogenetic tests are ordered by insurance companies, horse breeders, and owners to evaluate potential breeding stallions, broodmares, or the horses to be purchased. A more recent addition to equine clinical cytogenetics is the evaluation of chromosomal stability in stem cell lines that are produced for stem cell therapy.
Chromosomal analysis includes counting the diploid (2n) chromosome number, arranging chromosomes into a karyotype, and evaluating chromosome morphology according to their banding patterns following the international system of chromosomal nomenclature for the horse (ISCNH) (Bowling et al. 1997). The analysis also determines whether or not the chromosomal (genetic) sex agrees with the gonadal and phenotypic sex.
This chapter describes the methods that are used for clinical cytogenetic analysis in horses. This includes a list of the basic required equipment and supplies for an equine cytogenetics laboratory, a detailed step‐wise methodology for the preparation of metaphase chromosome spreads from different tissues, and a description of the main principles and methods of chromosome analysis. Finally, advanced chromosome analysis approaches and future perspectives of equine clinical cytogenetics are briefly discussed.
Chromosomal Analysis Techniques
Sample collection and shipping
Note: the underlying principle of chromosome analysis is cell division during which metaphase chromosome spreads can be obtained. Therefore, the material (blood, tissue, biopsy, etc.) for chromosome analysis must contain live cells and be free of bacterial and fungal contamination. This is why chromosome analysis cannot be done from hair or frozen tissues.
Peripheral blood (~20 ml) is collected in green top (for cell cultures) and purple top (for DNA isolation) Vacutainer® tubes, wrapped individually in paper towels to prevent breakage, and shipped with refrigerator packs (do not freeze!). Shipping should be overnight or as fast as possible. Blood older than 7–10 days may not be suitable for cell cultures. Equipment and Supplies Laboratory equipment Laminar flow hood for sterile cell culture work, CO2 incubator regulated at 37°C (99°F), clinical centrifuge with a swing‐out rotor for 15 ml conical centrifuge tubes, water pump for liquid aspiration, water bath, a set of adjustable volume laboratory micropipettes, autoclave, refrigerator‐freezer, heating plate or hot oven, inverted light microscope with 20× and 40× phase‐contrast objectives, light microscope with a 20× phase‐contrast objective and a 100× objective, equipped with a CCD (charge‐coupled device) camera and image analysis software to capture metaphase spreads and arrange karyotypes.Supplies for cell cultures 7–10 ml sodium or lithium heparin (green top) and EDTA (purple top) Vacutainer® tubes and 20 gauge needles for Vacutainer® tubes for blood collection, sterile forceps and small scissors, sterile Petri dishes, sterile T25 and T75 cell culture flasks, sterile conical 15 ml centrifuge tubes, sterile serologic pipettes(1, 2, 5, and 10 ml), culture medium for blood lymphocytes: RPMI medium 1640 with Glutamax™ and 25 mM HEPES buffer (Gibco™; Life Technologies Ltd, Grand Island, NY, USA) supplemented with 30% fetal bovine serum (FBS; various suppliers), 1× antibiotic–antimycotic solution (100× stock; Gibco), 1% pokeweed mitogen (lectin from Phytolacca americana; Sigma Aldrich, Burlington, MA, USA). (Larger quantities (500 ml) of ready‐to‐use media can be prepared and stored in 9 ml aliquots in sterile 15 ml screw‐cap centrifuge tubes at –20°C. The tubes should be defrosted only once.) Culture medium for fibroblasts: minimum essential medium (MEM) α with Glutamax™ (Gibco) and nucleosides supplemented with 10–20% FBS and 1× antibiotic–antimycotic solution, sterile Hanks’ balanced salt solution (HBSS; Gibco), sterile 0.25% trypsin‐EDTA (Gibco).Supplies for cell harvest and chromosome preparations Ethidium bromide 1 mg/ml solution in double distilled water (ddH2O) (optional; prevents chromosome condensation), demecolcine solution to stop the movement of cell spindle and capture chromosomes at metaphase (10 mg/ml in HBSS (Sigma‐Aldrich, St Louis, MO, USA)), hypotonic solution to burst cytoplasm and release the chromosomes (optimal hypotonic solution (Genial Helix, Chester, UK) or 0.067 M KCl solution in ddH2O), fixative: methanol/glacial acetic acid in a ratio of 3:1, pre‐cleaned double‐frosted microscope slides, fine‐tip plastic transfer pipettes, Coplin jars.Supplies for chromosome staining and banding Giemsa stain (KaryoMAX® Giemsa Satin Solution; Gibco), Gurr buffer tablets (Gibco), 0.125% trypsin (stock 2.5%; Gibco), 0.1N HCl, 5% Ba(OH)2 in water, 2× saline sodium citrate (SSC) (stock 20× SSC).
A skin biopsy is collected for transport into sterile tubes containing collection medium: 10 ml HBSS supplemented with 2× antibiotic–antimycotic solution.
Cell lines: pre‐established cell lines can be shipped frozen in cryovials on dry ice or in culture flasks at room temperature. Culture flasks must be completely filled with medium to prevent detachment and damage of cells during transport.
Peripheral blood lymphocyte cultures
Upon arrival, blood samples should stand at room temperature for 30 minutes to sediment the red blood cells. The buffy coat, with lymphocytes, is seen as a thin white layer between the red blood cells and plasma.
Under sterile conditions, add 1 ml of plasma with the buffy coat to pre‐warmed (37°C) 9 ml cell culture media in a 15 ml centrifuge tube. Mix the tube and incubate for 72 hours at 37°C. Gently invert the cultures twice a day. Note: only blood collected in Na‐ or Li‐heparin Vacutainer® tubes is suitable for cell cultures.
Start cell harvest at 68 hours of culture following the procedures described under “Cell harvest.”
Primary fibroblast cultures
Place a tissue biopsy, with a small amount of collection medium, into a sterile Petri dish and mince with sterile forceps and scissors to the size of 2–4 mm2. Wash the pieces 10 times in sterile collection medium (use a succession of Petri dishes).
Transfer the pieces of tissue into T25 flasks, approximately 4–6 pieces per flask. Add 0.5 ml culture medium and incubate at 37°C with 5% CO2 overnight. The pieces should get attached on the next day.
Check the pieces under an inverted microscope every day for the outgrowth of fibroblast cells, as well as for possible contamination. In the latter case, the cultures should be discarded.
Wash the pieces with HBSS/2× antibiotics–antimycotics and change medium (0.5 ml) every day. Do not let the pieces dry out. The first cells typically appear in a week.
When outgrowth is sufficient (~100 fibroblast cells around most of the pieces), remove the tissue pieces, aspirate the culture medium and wash the flasks twice with 5 ml of HBSS. Add 1 ml 0.25% trypsin‐EDTA and incubate for 5–10 minutes at 37°C to detach the cells. Without removing trypsin, add 2 ml culture medium, transfer the cells into a new T25 flask, add 5 ml fresh medium, and incubate at 37°C in 5% CO2. Check the cultures every day and change media every 2–3 days.
The cells can be grown in T25 flasks or, if more cells are needed, in T75 flasks. The transfer of cells from T25 to T75 is done with 0.25% trypsin‐EDTA as described in the previous step.
Cultures are ready for harvest when they are semiconfluent (~60–70%) and contain many mitotic cells visible as round, enlarged, partially detached bodies. Non‐diving fibroblasts appear as elongated bodies attached to the surface (Figure 22.1).
Note: Even though procurement and handling of biopsies and setting up primary fibroblast cultures is more laborious and time consuming compared with lymphocyte cultures, the advantage is that live fibroblasts can be cryopreserved in liquid nitrogen and used years later for additional cytogenetic studies and various advanced