Chapter 14 - Springer Link

Chapter 14 Tissue Fixation and Processing for the Histological Identification of Lipids Víctor Carriel, Fernando Campos, José Aneiros-Fernández, and John A. Kiernan Abstract Lipids are a heterogeneous group of substances characterized by their solubility in organic solvents and insolubility in water. Lipids can be found as normal components of different tissues and organs, and they can be affected by several pathological conditions. The histochemical identification of lipids plays an important role in histopathological diagnosis and research, but successful staining depends on adequate fixation and processing of the tissue. Here we describe methods to fix and process tissue samples for the histochemical identification of lipids in frozen or paraffin-embedded tissues. Key words Lipids fixation, Formaldehyde, Freezing techniques, Sucrose cryoprotection, Isopentane, Liquid nitrogen, Paraffin embedding, Linoleic acid, Lecithin, Lipid histochemistry

1 Introduction Lipids vary greatly from the structural and molecular point of view. They are soluble in organic solvents and insoluble in water [1]. Histochemically, lipids can be classified according to their hydrophobicity and hydrophilicity, or based on the presence or absence of esters and amides [1]. Lipids are essential for the normal function of the organism, playing important structural, metabolic, or endocrine roles. In this regard, significant amounts of lipids are present in adipose tissue, in sebaceous and adrenal glands [2], and in the myelin sheaths of central and peripheral nerve fibers [3, 4]. Lipids can be affected by several pathological conditions [5]. In this context, the histochemical identification of lipids plays an important role in histopathological diagnosis (e.g., atherosclerotic lesions, liposarcoma and demyelinating diseases) [2, 5] as well as in research. The aim of this chapter is to describe procedures for tissue fixation and processing for subsequent lipid histochemistry in

Carlo Pellicciari and Marco Biggiogera (eds.), Histochemistry of Single Molecules: Methods and Protocols, Methods in Molecular Biology, vol. 1560, DOI 10.1007/978-1-4939-6788-9_14, © Springer Science+Business Media LLC 2017

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sections of frozen or paraffin-embedded material. The starting point of each method is chemical fixation of the tissue. Fixation is the first and most critical step in histology [1, 6, 7], intended to immobilize the cellular and extracellular material in a condition closely resembling the structure and chemical composition of the living tissue [1, 6, 8]. Specimens can be fixed by chemical reactions, physical methods (such as heating and freezing), or by combination of both methods [1, 6, 7]. Fortunately, most of lipids are unaffected by the fixation with non-coagulant chemical agents, notably formaldehyde, which stabilizes structure by cross- linking protein molecules [1, 7]. Lipids are also well fixed (and also stained) by osmium tetroxide (for the method, see the Chapter 15) [9, 10]. Lipids are neither displaced not chemically changed by freezing [1, 2, 7]. In contrast, most lipids are extracted from tissues by organic solvent-based fixatives, which immobilize proteins by coagulation. The incorporation of calcium ions into an aqueous solution of formaldehyde (Baker’s formal-calcium) increases the preservation of hydrophilic phospholipids [1]. Our recommendation is for chemical fixation of specimens in 4 % neutral buffered formaldehyde (10× dilution of formalin, which contains 40 g formaldehyde per 100 ml) or formal-calcium. Immersion for 24–48 h is necessary for adequate cross-linking of proteins. Once fixed, samples can be frozen and sectioned. Alternatively, the specimens may be dehydrated, cleared and embedded in paraffin. The organic solvents used for dehydration and clearing extract most lipids; only lipids that are covalently bound to protein can be detected in routinely prepared paraffin sections. A post-fixation treatment described by Tracy and Walia [11, 12], however, renders lipids insoluble in organic solvents and allows their histochemical detection in paraffin sections. Currently, most lipid histochemistry is done on fresh (unfixed) frozen tissues, which are useful for the examination of biopsies for rapid diagnosis, and also for enzyme activity histochemistry and immunofluorescent techniques [1, 7, 13, 14]. However, it is often not feasible to obtain ideal conditions for rapid freezing. Ice crystal formation due to slow freezing produces irreversible damage to the tissue architecture (Figs. 1 and 2) [1, 7, 13, 14]. In order to improve tissue morphology and reduce damage due to ice crystal formation, biopsies must be first fixed in formaldehyde, then cryoprotected, and then frozen quickly [1, 14]. In this sense, tissues must be frozen by using cooled isopentane at −80 °C (freezer) or −160 °C (liquid nitrogen). This procedure minimizes ice crystal formation and allows the cutting of frozen sections of good quality, suitable for most lipid histochemical methods (Figs. 1 and 2) and other staining techniques. Conventional paraffin embedding provides internal and external mechanical support to the tissues for microtomy [8]. In the usual method the specimens are completely dehydrated by replacing

Tissue Fixation and Processing for the Histological Identification of Lipids

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Fig. 1 Examples of lipid histochemistry of adipose tissue carried out with described methods. Positive histochemical reactions with Sudan IV (a, c, and d) and oil red O (b) in adipose-rich tissues. Images a–c show results obtained in chemically fixed, cryoprotected and quickly frozen tissues, whereas image d shows the typical artifact (holes due to ice crystal formation) caused by slow freezing (−20 °C) of unfixed material. Image e shows complete extraction of lipids by organic solvents during the conventional paraffin embedding procedure

the tissue’s water with alcohol, “cleared” in an alcohol-miscible paraffin solvent such as xylene, and finally infiltrated with molten paraffin wax [1, 6, 7]. Although lipids are not affected by formaldehyde fixation, most are extracted from the tissue by the alcohol and xylene used during tissue processing. Some lipoproteins and phospholipids resist extraction during this process, and these can be stained by some histochemical techniques. This mainly occurs with the myelin sheaths of the central and peripheral nervous systems [3, 4, 6] (Fig. 2) (see staining methods on Chapter 15) [9]. Tracy and Walia [11, 12] have described a method that successfully preserves lipids in paraffin sections. The formaldehyde- fixed specimens are infiltrated with an emulsion of unsaturated lipids in ethylene glycol. These lipids partition into tissue lipids. A subsequent treatment with chromic acid changes the introduced lipids into organometallic complexes that are insoluble in organic solvents and stainable in paraffin sections. Finally, this chapter offers different procedures after formaldehyde fixation to perform general lipid histochemistry. The selection of one of these methods should be determined by the aims of the study. Tissue sections from frozen or paraffin-embedded samples can be used for other histochemical and immunohistochemical applications. The Tracy and Walia method was designed to allow

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Fig. 2 Images showing positive histochemical reactions of myelin in the central and peripheral nervous system. Sudan Black B (a, b, d, f, g), oil red O (c, e) and MCOLL (h) histochemical methods. Images a–f were obtained from tissues subjected to formaldehyde fixation, cryoprotection and fast freezing, while the image g shows damage due to the slow freezing (−20 °C) of a previously fixed, but not cryoprotected nerve. The image h shows myelin stained blue by the MCOLL method in a normal peripheral nerve fixed with formaldehyde and embedded in paraffin (for details of MCOLL method, see Chapter 15) [9]. Note the typical undulating course of nerve fibers (f–h) in longitudinal sections. If straight nerve fibers are desired, the fresh nerve can be stretched, under a dissecting microscope, until the bands of Fontana (transverse striations) just disappear. The ends of the specimen can then be pinned to a cork, or the stretched nerve can be placed on a small card for about 1 min, until it adheres, before immersing in the fixative

lipid histochemistry with paraffin sections. It is not yet known whether sections of chromated tissue are suitable for other histochemical or immunohistochemical techniques.

2 Materials In preparing specimens for lipid histochemistry workers will be in contact with fresh biological material and they will need to use some toxic reagents. Toxic solutions must be prepared under a laminar flow hood using lab coat, gloves, and, when necessary, laboratory goggles and a mask. Tissues must be handled, fixed, and dissected in a pathology workstation (ideally) or in a chemical fume hood, while wearing


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medium or heavyweight nitrile, neoprene or PVC gloves, tight-fitting safety goggles or a full face shield. Similarly, for freezing techniques, adequate gloves and eye protection are needed. Isopentane is flammable and volatile (boils at 28 °C). 2.1 Materials and Equipment for Tissue Fixation

1. Biopsy containers or similar. 2. Magnetic stirrer. 3. Dissecting instruments. 4. Electrothermal blowing drying oven or hybridization oven at 56 °C. 5. Separatory funnel.

2.2 Fixation Reagents and Solutions

1. Distilled water. 2. 0.1 M phosphate buffer solution (PB), pH 7.2–7.4. 3. 4 % neutral buffered formaldehyde solution. Dilute formalin (40 % HCHO, w/v) to 10× volume by adding PB. This solution can be used immediately. 4. Formal-calcium (Baker’s formal-calcium) solution. Dilute formal dehyde (37–40 % HCHO, w/v) to 10× volume with a 2 % w/v aqueous solution of calcium acetate (monohydrate) [1]. 5. 70 % v/v solution of ethylene glycol (analytical reagent grade) in water. 6. 70 % v/v solution of ethanol (analytical grade) in water. 7. Linoleic acid, technical grade (60 %; the other major component of this liquid is oleic acid). Alternatively use pure (99 %) linoleic acid. 8. Lecithin (product sold as a dietary supplement is suitable). 9. Solution 1. Add either 20 ml of technical grade linoleic acid [12] or 5 g of pure (99 %) linoleic acid [11] and 2 g lecithin to 500 ml 70 % ethylene glycol. Mix for 4 h on a magnetic stirrer and then leave it to stand for at least 4 h. Finally, draw off the lower phase in a separatory funnel. This emulsion is stable for several weeks, but the separation may need to be repeated if an oil layer forms on the surface. 10. Solution 2. 2 % aqueous solution of chromium trioxide (“chromic acid”). 11. Solution 3. 5 % aqueous solution of sodium bicarbonate (analytical reagent grade).

2.3 Freezing Technique Reagents and Equipment

1. Laboratory freezer (−80 °C). 2. Liquid nitrogen, tank and pump. 3. Isopentane (2-methylbutane). 4. 20 % aqueous solution of sucrose.

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5. Falcon tubes or similar (between 15 and 50 ml). 6. Freezable plastic or metal bottle with wide cap (for isopentane). 7. Cryomolds, metallic holders or similar. 8. Cryoprotective agent OCT (optimal cutting temperature) or similar. 9. Polystyrene box with wide cap (for liquid nitrogen). 10. Large forceps (to handle samples during freezing). 11. Sealable plastic bags. 2.4 Tissue Processing, Paraffin Embedding, and Equipment

1. Histological cassettes. 2. Glass or xylene-resistant plastic vials (at least 12) with wide caps, or an automatic tissue processor. 3. Electrothermal blowing drying oven at 60 °C for paraffin or commercial embedding center. 4. Analytical grade ethanol solutions (50 %, 70 %, 95 %, 99 %). Ninety-nine percentage is often sold as “absolute” or “anhydrous” ethanol; it can be diluted with distilled water to make the lower percentages. 5. Xylene (or an alternative clearing agent such as d-limonene). 6. Paraffin wax for histology (see Note 1). 7. Paraffin embedding molds.

3 Methods Lipid histochemistry can be performed with frozen or paraffin- embedded tissues. In all cases, adequate fixation is essential for all subsequent histological techniques. Biopsies must be fixed as soon as possible in an adequate volume of fixative. It is important to follow precautionary measures when working with formaldehyde. If fixative is not available, samples can be transported to the laboratory (as soon as possible) in a biopsy container with wet gauze (saline solution) at 4 °C in a polystyrene box. Once in the laboratory, samples must be properly labeled. We recommend that you prepare all materials, solutions, and equipment in advance. 3.1 Fixation

1. A whole organ or other large specimens must first be immersed in at least ten times its own volume of either neutral buffered formaldehyde or formal-calcium, for a few hours (1–2 h). 2. When organs or biopsies have hardened somewhat, cut them and select representative samples no more than 2 mm thick, and place them in histological cassettes or Falcon tubes (see Note 2). 3. Complete fixation by immersing cassettes or samples in at least ten times their volume of neutral buffered formaldehyde or formal-calcium.


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4. Leave the samples in the fixative for 24–48 h at room temperature, with agitation (see Note 3). 5. Rinse fixed samples in tap water and then proceed with free zing or conventional paraffin embedding or with Tracy and Walia lipid post-fixation method. 3.2 Freezing Technique

1. Immerse fixed and rinsed samples in ten times the sample volume of 20 % sucrose solution for 24 h at 4 °C. 2. Cool a sufficient volume of isopentane in a freezable bottle with wide cap at −80 °C (−80 °C freezer) or −160 °C (liquid nitrogen) (see Note 4). 3. Place cryoprotected sample (or fresh material) in a cryomold and cover it completely with OCT. 4. Hold the samples with a large forceps and freeze by immersion on isopentane at −80 °C (in a −80 °C freezer) or at −160 °C (in isopentane immersed in liquid nitrogen) (see Note 5). 5. Mount frozen samples into the cryostat specimen chuck, and cut sections 10–15 μm thick (for storage of samples, see Note 6).

3.3 Paraffin Embedding

This method is suitable for formaldehyde-fixed samples (routine method) and samples post-fixed with Tracy and Walia method. The routine method allows us to stain myelin and some lipids that resist extraction. In contrast, Tracy and Walia’s method improves the preservation of all lipids, including neutral fat and cholesterol. Here we describe standard schedules for routine and rapid tissue processing, but the times can be adapted according to sizes and types of samples [1, 8]. In addition, throughout the procedure, the volume of liquid should be 10–20 times that of the specimen [1] (for general recommendations, see Note 7).

Reagent

Routine (overnight)

Rapid (small samples), min

1. 70 % ethanol

1 h

25

2. 95 % ethanol (1)

1 h

15

3. 95 % ethanol (2)

1 h

15

4. 95 % ethanol (3)

1 h

30

5. 99 % ethanol (1)

1 h

15

6. 99 % ethanol (2)

1 h

15

7. 99 % ethanol (3)

1 h

30

8. Xylene or similar (1)

30 min

10


45 min

15


45 min

15 (continued)

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Reagent

Routine (overnight)

Rapid (small samples), min

11. Paraffin wax (1 at 60 °C)

1 h

30


1 h

30


1 h

30

3.4 Tracy and Walia Lipid Post-fixation Method

Note that this method requires 5–7 days, but only needs few minutes of active effort in this period of time. This method is suitable for freezing technique, but an important improvement of lipid histochemistry can be achieved in formaldehyde-fixed and paraffin- embedded tissues [11, 12]. 1. Immerse formaldehyde-fixed tissues (no more than 2 mm thick) in solution 1 for 3 days at 56 °C (see Note 8). 2. Rinse samples in several changes of 70 % ethanol for a minimum of 8 h. 3. Rinse in several changes of water for 8 h. 4. Immerse in solution 2 for 24 h at 4 °C. 5. Rinse in tap water for 24 h at room temperature. 6. Immerse in solution 3 for 24 h at room temperature. 7. Rinse in tap water for at least 8 h. 8. Proceed with the dehydration, clearing and paraffin embedding as described in Subheading 3.3 above.

4 Notes 1. Melting point of paraffin waxes range from 52 to 58 °C and as a general rule, a higher melting point gives a harder block. Waxes sold for histology commonly contain added polymers, not identified by the manufacturers but claimed to accelerate the penetration and increase the hardness. It is important to keep wax about 2 °C above its melting point. Higher temperatures are said to damage the secret additives. 2. It is important to perform a correct tissue dissection and to obtain thin representative samples that are no more than 2 mm thick. Thicker samples will increase the times needed for all steps (fixation, freezing, dehydration, clearing and paraffin embedding). In addition, for the Tracy and Walia method thin tissue blocks are necessary for uniform penetration of the post- fixative solutions. 3. The duration of fixation depend of several factors (sample size, type of tissue, temperature, static vs agitation conditions, etc.). Formaldehyde can correctly fix a small object in 8 h, but we


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recommend you fix specimens for at least 24 h. Large samples and/or dense tissues will need longer times. In the case of neural tissue (brain and spinal cord) it is recommended to fix samples for at least 48 h or even longer. 4. Isopentane must be stored in an explosion-proof refrigerator at 4 °C or less, because it is an extremely volatile and flammable liquid with high vapour pressure at room temperature. It is necessary to use lab coat, gloves and mask. Isopentane can be cooled at −80 °C 1 or 2 days before using; it will not freeze at this temperature. To cool isopentane to its freezing point (−160 °C) it is necessary to deposit enough liquid nitrogen in a polystyrene box and then immerse the bottle with the isopentane for some minutes. The isopentane becomes viscous, then gelatinous, before it freezes. 5. To freeze the sample at −80 °C by using a cooled isopentane and −80 °C refrigerator, immerse the cryomold with the sample immersed in OCT directly into the isopentane for 3–5 min. To perform a quick freezing it is necessary to immerse the samples into cooled isopentane in liquid nitrogen until OCT is completely frozen. Long exposure to liquid nitrogen should be avoided because it could cause tension cracks. 6. The −80 and −160 °C frozen samples can be packed in a plastic bag, sealed and stored at either −25 or −80 °C for at least 5 months. 7. In general, tissue processing should be performed manually in a chemical fume hood or by using automatic tissue processor. For manual processing, the worker should wear lab coat, gloves, and mask, especially during fixation and clearing. Deli cate tissues must start dehydration from 30 or 50 % ethanol solutions (or even lower) [1]. An example of a delicate object is a peripheral nerve sample subjected to the osmium tetroxide technique (see Chapter 15). Times are standard for general purposes, but it is important to take into account that the size and type of tissue will have an impact on this process. In general, tissue blocks of 1 mm thickness require around 30 min in each reagent, while a tissue blocks of 5 mm thickness could require around 90 min or more in each reagent. An automatic tissue processor can be programmed to work overnight, and it is optional to start with 70 % ethanol or 10 % neutral buffered formaldehyde (especially if it is necessary to complete the tissue fixation), but this will leave traces of formaldehyde in the alcohols. Manual processing needs stand points, especially for long procedures. Specimens may be left overnight in 70 or 95 % ethanol at 4 °C, but not in 99 % ethanol or xylene, which can cause excessive hardening of tissues).

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8. For the Tracy and Walia method it is recommended to processes all the samples by using glass bottles. Histological cassettes are not recommended, because they affect the penetration of solutions into the tissues.

Acknowledgments The methods described in this chapter were supported by the Spanish “Plan Nacional de Investigación Científica, Desarrollo e Innovación Tecnológica”, from the National Ministry of Economy and Competitiveness (Instituto de Salud Carlos III), Grant No FIS PI14-1343 and cofinanced by “Fondo Europeo de Desarrollo Regional” (FEDER), European Union; The Tissue Engineering Group (CTS-115) of the University of Granada, Spain; and by the “Fundación Benéfica Anticáncer San Francisco Javier y Santa Cándida, Granada, España”. References 1. Kiernan JA (2015) Histological and histochemical methods: theory and practice, 5th edn. Scion, Banbury 2. Mills SE (2007) Histology for pathologists, 3rd edn. Lippincott Williams & Wilkins, Philadelphia, PA 3. Carriel V, Garzón I, Alaminos M et al (2011) Evaluation of myelin sheath and collagen reorganization pattern in a model of peripheral nerve regeneration using an integrated his tochemical approach. Histochem Cell Biol 136(6):709–717 4. Kiernan JA (2007) Histochemistry of staining methods for normal and degenerating myelin in the central and peripheral nervous systems. J Histotechnol 30(2):87–106 5. Kumar V, Abbas AK, Aster J (2013) Robbins basic pathology, 9th edn. Elsevier Saunders, Philadelphia, PA 6. Carriel V, Garzón I, Alaminos M et al (2014) Histological assessment in peripheral nerve tissue engineering. Neural Regen Res 9(18): 1657–1660 7. Cook DJ, Warren PJ (2015) Cellular pathology: introduction to techniques and applications, 3rd edn. Scion, Bloxham

8. Crocker J, Burnett D (2005) The science of laboratory diagnosis, 2nd edn. John Wiley, Chichester 9. Carriel V, Campor A, Alaminos M, Raimondo S, Geuna S (2017) Staining methods for normal and regenerative myelin in the nervous system. Methods Mol Biol Volumen 1560, Histochemistry of single molecule, methods and protocols. ISBN: 978-1-4939-6787-2 10. Di Scipio F, Raimondo S, Tos P et al (2008) A simple protocol for paraffin-embedded myelin sheath staining with osmium tetroxide for light microscope observation. Microsc Res Tech 71(7):497–502 11. Tracy RE, Walia P (2002) A method to fix lipids for staining fat embolism in paraffin sections. Histopathology 41(1):75–9 12. Tracy RE, Walia P (2004) Lipid fixation for fat staining in paraffin sections applied to lesions of atherosclerosis. Virchows Arch 445(1):22–26 13. Meier-Ruge WA, Bruder E (2005) Pathology of chronic constipation in pediatric and adult coloproctology. Pathobiology 72(1-2):1–102 14. Serrato D, Nieto-Aguilar R, Garzón I et al (2009) Comparison of the effect of cryopreservation protocols on the histology of bioengineered tissues. Histol Histopathol 24(12):1531–1540

Chapter 14 - Springer Link

Chapter 14 - Springer Link

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