Histological and cell biological methods
1-3. Vital stainings, sample preparation methods and fixation, Freezing-microtomy, enzyme histochemistry (acid and alkaline phosphatase, esterase, lipid and others) Embedding in paraffin, sectioning and staining for routine histology (toluidine blue, hematoxilin-eosin, PAS and others)
4. Confocal microscopy Introduction: the confocal laser microscope and the semi-confocal fluorescent microscope (spinning disc, apotome), the multichannel image recording and the combination of images. The combined use of various fluorescent dies. Practical: native and fixed samples, vital staining of organelles, GFP-expression, multiple labelling and colocalisation. Photography and serial recordings, optical slicing, 3D and !D images. Computer processing of images and evaluation.
5.-6.-7. Electron microscopy Introduction: Principles of electron optical imaging, electronic lenses, diffraction, resolution. Interactions of the material with the electrons and their use for image generation and chemical analysis. The technical solutions in electron microscopy, transmission and scanning microscopes. Lens imperfections and their correction, apertures. Enhancement of contrast, image collection and storage. Sample preparation: fixation, embedding, sectioning and staining. Evaluation of the pictures. Practical: chemical fixation by aldehydes and osmic acid, cryofixation and cryosubstitution. Embedding, ultramicrotomes, making glass knifes,cutting, contrasting, negative staining. Demonstrations of the performance of the transmission electron microscope: electron optical adjustments, checking resolution, observation of samples and making photoghraphs.Vacuum evaporation techniques.
8-9. Image processing and quantitative morphology Image fixation and collection, analogue and digital images, their advantages and disadvantages. File format and patents, making archives. The mathematical and computer bases of digital image transformations and manipulations. Qualitative and quantitative data and measurements in practice.
10-11. Cell fractionation Introduction: the principles of cell fractionation and centrifugation methods, measurement of the density of sedimenting particles, precipitation time,its estimation and measurement. Differential centrifugation, continuous and discontinuous density gradients, zonal methods. The theoretical ground of homogenization. Practical: homogenization. Fractionation by differential centrifugation, raw and refined fractions, gradient centrifugation, self creating Percoll gradient. The utilization of markers in checking the purity of fractions.
12-14 In vitro methods Perfusion techniques, basic technical parts and types, applicability, advantages and disadvantages application conditions. In situ, isolated, single-pass and recirculation systems, perfusion media. Embryonic and adult organs in simple surviving cultures, organotypic and histiotypic growth. Conditions of survival, creation and experimental utilization of non-perfusion organ cultures. In vitro fertilization and whole embryo cultures. Real tissue and cell cultures, and their utilization. Technical ground and working conditions (equipment, sterility, media, gas phase, safety rules). Explant and isolation techniques, primary cultures, cell lines, cell strains, clones, long-term storage. Practical approach of various experimental problems. Subcellular in vitro systems. Practical demonstrations, digital and video material.
15. The application of reporter-genes in cell biology Introduction: the reporter-gene, the most widely used reporters and their applications. Preparation of reporter constructs and their transfer into the cell or organism, transgenic organisms. Practical: Gyakorlati rész: observations on two lacZ transgenic Drosophila strain and the wild type: L3 wandering larvae, dissection, fixation, X-gal staining. GFP-tagged proteins and their use in functional and expression analysis.
Hand-out printed materials from the lecturer