Bone is considered the most important supporting tissue in the body. It is composed of cells, organic extracellular matrix and inorganic salts. Bone tissue is mineralized in layers which provide great strength and flexibility to the skeletal system. It varies in formation, depending on its function, across the body. These functional/formation differences are also based on the proportion of the different inorganic and organic processes incorporated or produced in the formation of a bone. The most common mineral in bone is hydroxyapatite which consists of collagen, proteins and carbonate ions. The main bulk of bone is approximately 70% mineral and 30% organic components by weight. Bone cells, as opposed to marrow cells, are relatively sparse. This chapter will review bone morphology and its organic and inorganic components, as well as considering methods on preparing sections of bone for analysis that can be used in clinical and research histology laboratories.
Two types of bone can be recognized macroscopically in the adult human skeleton. They are cortical or compact bone and trabecular, cancellous, or spongy bone. Compact bone is the solid, hard, and immensely strong bone that forms the shafts of long bones (e.g. femur and tibia) and exterior surfaces of the flat bones (e.g. ribs and skull). Trabecular bone is found in the diaphysis, epiphysis, and marrow cavities of long bones, vertebrae, and centers of flat bones. It is a mesh of bone strands each about 1 mm thick. Although it looks less solid than cortical bone, this arrangement of trabeculae, particularly in the femoral head and vertebrae where it forms an almost ideal weight-bearing structure, is very strong.
The major components of bone are mineral, cells and an organic extracellular matrix of collagen fibers and ground substance. These are dynamic components as the processes of cell replacement, repair and remodeling of bone, and the erosion and reformation of collagen and mineral occur continually throughout adult life.
The collagen found in bone differs from other collagen in the body in that it becomes mineralized and is laid down in bands or lamellae roughly parallel to one another. The collagen fibers within each lamella tend to lie next each other but at an angle to the fibers in adjacent lamellae. A cement of proteoglycan ground substance outlining these fibers is seen in sections only at the cement lines. The organization of collagen lamellae is responsible for the distinctive micro-anatomical patterns of bone which are easily seen with polarized light microscopy (Fig. 16.1).
The simplest pattern occurs on the periosteal and endosteal surfaces of compact bone as circumferential lamellae, and in trabecular or non-Haversian bone where lamellae are roughly parallel to the surface. Cortical bone is composed of Haversian systems or osteons in which concentric lamellae surround channels (Volkmann’s canals) containing one or more blood vessels. These tubular structures run longitudinally in the bone and are packed closely together with irregular interstices filled by the remnants of older osteons (Fig. 16.2). Cement lines outline the boundaries of osteons, and some trabecular and circumferential lamellae.
Figure 16.2 Haversian systems (osteons, Volkmann’s canals, and bone cells) in ground section of undecalcified bone in methyl methacrylate, acid etched/surfaced stained with McNeal’s/toluidine blue stain.
Another collagen fiber arrangement forms non-lamellar or woven bone and is found in immature bone and some pathological conditions. This collagen is not deposited in the lamellae but in thick, short, randomly oriented bundles. When viewed with polarized light, these appear as coarse fibers resembling a woven fabric.
Unmineralized collagen or osteoid forms a border, or seam, on surfaces of newly formed bone, and after osteoid is deposited there is a lag before it becomes mineralized. Osteoid is normally around 15 µm thick, covering only a small proportion of the surfaces. In some diseases, such as rickets or osteomalacia, it is much thicker and widespread.
On inactive surfaces the osteoid is very thin, difficult to see, and is completely absent where resorption is taking place. This process is called remodeling and consists of resorption and deposition taking place in equilibrium so that the volume and shape of bones stays more or less constant. In later life, remodeling slows down, and deposition may not keep up with resorption, causing increased bone porosity and brittleness, and, in extreme cases, the disease osteoporosis.
The main mineral content of bone is calcium and phosphate combined with hydroxyl ions to form hydroxyapatite crystals. The mineral is approximately 38% calcium and thought to be deposited as amorphous calcium phosphate in the initial mineralization phase. This transforms to hydroxyapatite by addition of hydroxyl ions to form a crystal lattice into which carbonate, citrate, and fluoride ions as well as magnesium, potassium, and strontium can be substituted or included. Carbonate is present in large quantities but probably only in the hydration shell and on crystal surfaces.
The hydroxyapatite forms needle-like crystals about 22 nm in length, resulting in an enormous total crystal surface area. Thus, the mineral fulfills the obvious function of giving strength and rigidity while approximately 20% remains in the amorphous form to provide a readily available buffer for maintaining total body chemical equilibrium (e.g. pH and enzyme system regulation).
Osteoblasts are fully differentiated to carry out the primary function of bone formation by producing and laying down osteoid. They are seen on surfaces of actively forming bone as plump cells with basophilic cytoplasm and eccentric nuclei distal to the bone surface. The cytoplasm is basophilic due to ribonucleic acid, and before becoming fully differentiated frequently contains glycogen. Acid phosphatase is found in osteoblasts and the surrounding tissues, but decreases at onset of calcification. Upon completion of bone formation, most osteoblasts revert to a quiescent (small undifferentiated cell) state and reside among the heterogeneous cell population.
In one respect osteoblasts do represent immature cells, as some are trapped in the osteoid matrix they lay down, and become mature osteocytes residing in tiny spaces or lacunae within the bone. Lacunae are connected to each other and to the vascular spaces by canaliculi – tiny channels into which osteocyte processes project for the purpose of passing fluids and dissolved substances necessary for cell metabolism (Fig. 16.3).
These are the cells responsible for bone resorption or erosion. Osteoclasts are large multinucleated (giant) cells whose cytoplasm contains numerous mitochondria and alkaline phosphatase. They occur in small clusters or singly on bone surfaces undergoing resorption and are often seen in the depressions (Howship’s lacunae) they are actively creating by erosion. These surfaces have an irregular outline and lack osteoid (Fig. 16.4). The direction of resorption is random, with no relationship to lamellar structure. Osteoclasts respond to altered mechanical stresses on the skeleton and to growth, and their activity contributes to remodeling. They also respond to hormones that can either stimulate or inhibit their activity. When resorption halts, the process of bone formation resumes (osteoid, mineralization, etc.). Cement lines will occur at the junction between old and newly formed bone.
This occurs in flat bones, e.g. skull, sternum, pelvic bones. A fibrous membrane first develops at the bone formation site in which mesenchymal cells differentiate into osteoblasts that begin the bone formation process by laying down osteoid. This starts in small islets that gradually unite to become trabeculated and finally form an external layer of compact bone.
This occurs in long bones and major parts of the skeleton. This type of bone development begins with differentiation of mesenchymal cells at sites where bone will be formed, but this bone is laid down in a cartilage model that resembles the final shape of the bone. This cartilage model becomes covered with a connective tissue sheath or perichondrium and grows by both apposition and interstitially. Appositional growth or the laying down of more cartilage begins towards the exterior and is mainly responsible for increased diameter; interstitial growth is by cells dividing within the model and mainly occurs towards the extremities, resulting in increased length.
In the central part of the model, cells continue to differentiate, cartilage begins to calcify, blood vessels invade, and the cartilage is broken up into strands. Ossification, by differentiation of perichondrial cells into osteoblasts, begins around the exterior of the primary ossification site at the center of the model. Osteoblasts invade the strands of calcified cartilage and deposit osteoid that soon becomes calcified. This process continues and secondary ossification sites appear at each end of the model, separated from the new bony shaft by cartilage growth or epiphyseal plates capable of interstitial growth. Once bone has ossified, it can only grow by apposition. Remodeling is continuous. In some places deposition exceeds resorption, whereas in others this is reversed and results in the characteristic shape of the bone. When bone is fully grown, the three ossification sites unite and the cartilage growth plates disappear.
The technique chosen for examination of bone is influenced by the initial clinical diagnosis, case urgency, and the extent of investigation required. Specimens arriving in the laboratory can vary in size from a needle biopsy a few millimeters long to whole appendages, e.g. amputation. Mineralized sections are used for microradiographic and histomorphometric studies as well as polarized and fluorescent light microscopy.
Biopsies are used for diagnosis of several diseases such as cancers, hemopoietic disorders, and infections. These specimens are usually small enough to treat much like soft tissue, except that a bone biopsy usually needs decalcification. This is particularly neccesary if it contains a piece of the cortical bone in order to produce paraffin sections. A bone marrow biopsy is usually removed with a Jamshidi needle for diagnosis of metabolic bone disease. Metabolic bone diseases are diagnosed using trabecular bone (Byers & Smith 1967) taken from the iliac crest which is an accessible bone site representative of skeletal bone as a whole.
Sections that are requisite to assess the relationship between mineralized and unmineralized bone (osteoid) are best processed and embedded into a plastic, such as methyl methacrylate (MMA), glycolmethacrylate or a epon-like plastic. The preferred method used for metabolic bone disease research and diagnosis is MMA plastic. Plastic is not a common embedding medium in clinical laboratories but is useful in research and clinical laboratories that are connected with research. It is also possible to produce frozen sections from an undecalcified bone biopsy. There are silver stains that demonstrate bone and osteoid in a decalcified, paraffin-embedded bone section (Tripp & McKay 1972) but many researchers choose the MMA-embedded section.
It is not practical to bisect (half for paraffin/half for plastics) an iliac crest trephine biopsy if both paraffin and plastic embedding methods are employed in the laboratory. Metabolic bone disease laboratories usually prefer a whole trephine bone core for plastic embedding. Needle biopsies should remain whole for paraffin or plastic methods.
Large amputation specimens are usually taken as a result of tumor, chronic osteomyelitis, and gangrene. These specimens are often delivered to the laboratory immediately after removal. They often are not in any sealed container and without fixative, and must be dealt with as soon as possible (either in the mortuary or laboratory). The majority of the limb is usually discarded or saved/fixed (if requested by patient) and the area or lesion with actual or suspected involvement in the disease process is retained for final evaluation. Skin, excess muscle, and connective tissue should be cut away from the lesion if possible. Excess bone or a joint disarticulated above and below the lesion should be performed so that fixation is adequate. The relevant portions should be immersed into a large volume of fixative to insure complete fixation. If it is not possible to inspect the specimen for several hours after receipt, it should be refrigerated at 4°C or it can be placed in fixative as a whole and kept at 4°C. Placing it in fixative prior to trimming helps in managing the trimming and prevents autolysis of the outer layers of the specimen. The mortuary/morgue area has a dual advantage for both limb storage and subsequent sample preparation on an available autopsy table. Whenever possible, specimen radiography of large bone specimens helps select the lesion/diseased area for trimming to a smaller sample size for processing.
Benign or low-grade malignant tumors and arthritic femoral heads resemble large biopsy specimens, frequently have an established diagnosis and are often considered less urgent. In femoral head or knee replacement surgeries, the bone specimen removed from the patient is usually received in the laboratory whole. Either prior to or after some fixation, a wedge-shaped sample is cut from the whole specimen using a Stryker bone saw or a heavy-duty X-ACTO knife. This wedge-shaped sample is placed back into fixative for 24–48 hours and then decalcified, processed and sectioned for pathology evaluation.
Unless immediate diagnosis is needed using cryomicrotomy, all bone specimens must be totally fixed before subjecting them to any decalcification and processing procedures. Complete fixation helps protect bone and surrounding soft tissue from the damaging effects of acid decalcification. Ten percent neutral buffered formalin (NBF) is suitable for both paraffin and non-tetracycline labeled bone. It should be noted that fixation proceeds faster by reducing the size of the bone, opening the bone, and removing excess skin and soft tissue surrounding the lesion. Large specimens can be bisected or reduced in size by sawing into multiple slabs, and immersed into fixative immediately, or no longer than 48 hours after initial fixation. Once cut into smaller pieces, the samples should be placed into fresh fixative.
For MMA embedding, 10% NBF is generally used for fixation. Alcoholic formalin or 70% ethanol fixation is the fixative of choice for tetracycline-labeled bone. Alcohol-based fixatives are not recommended for bone destined for acid decalcification as alcohol can slow or prevent decalcification. Fixatives containing chloroform (Carnoy’s) and mercury (B5, Zenker’s, Susa), including substitutes of them, should be avoided for specimens to be radiographed since those fixatives tend to make bone radio-opaque and unsuitable for specimen interpretation.
Good saws are an essential piece of equipment in a bone histology laboratory. Other than surgical saws, there are a range of hobby shop or handyman’s bench saws that are designed to cut through stones, plastic, and some thin metals. These saws cut through cortical bone slowly with cuts no deeper than 7.5 cm. Dry saws may need slight modifications to prevent blade slippage when cutting wet, fatty bone. Water-cooled saws prevent heat damage to bone due to high-speed sawing, and are capable of full-length cuts through long bones and appendages, e.g. femur and tibia. Buehler Isomet Low Speed Saws (Buehler Ltd, USA) are used for trimming specimens, plus cutting bones embedded in plastics. This type of saw has a thin diamond-impregnated blade with a water-cooling bath and is ideal for bisecting biopsies (only when required) and larger bones (depending on the bone specimen diameter). It can make precise, debris-free cuts through 8 mm thick bone cores, other cortical or trabecular and MMA-embedded bone specimens. Scalpel blades, fretted wire or jewelers’ saws have been used to cut biopsies with damaging results. These cutting devices can crush or fracture fine trabeculae, creating ‘fracture artifact’, and force bone fragments into marrow spaces, spoiling the bone histology.
Suitable blade specifications on small saws are 0.5 cm width and 12 to 16 teeth per inch (tpi), making finer, cleaner cuts than a larger saw blade 1.25 cm wide with 6 tpi. Blades in specifications needed are available from tool companies.
Soft tissues and dense connective tissue, e.g. tendons, should be removed before sawing or the sample will drag through the blade. The first cut is made through the mid-plane, then approximately 3–5 mm thick slabs are cut parallel to the first cut. A saw guide plate or wood block held against the first cut edge ensures an even slice. It is safer for workers to hold thinner bones between two wood blocks that will not ruin blades. Sawing should be at a slow even rate to match speed of blade cutting into the bone. Pushing the bone produces uneven cuts, and may jam or break a blade.
Bone slabs should be fixed for an additional 24–48 hours, especially if they appear pinkish-red or partially fixed. After sawing, any bone dust or debris adhering to slab surfaces can be cleaned away using a slow stream of water and a wet paper towel to brush off the debris. Care must be taken not to push debris into marrow spaces or to wash slabs excessively before the bone is totally fixed.
Thin bone slices give sharper-image radiographs than a whole specimen or clinical radiographs. When using film and not digital images, the use of ‘soft’ X-rays (of low kV) and high-contrast X-ray film provides finer detail and clarity (Fornasier 1975).
Many non-specialist departments of pathology use their in-house radiology department. However, there is the ‘Faxitron’ (Faxitron Inc., USA) cabinet X-ray system which can be used for specimen X-rays of bone in clinical and research settings. It comes in a free-standing and a tabletop unit. The energy output is 10–110 kV with 3 mA tube current. In addition to the manual exposure capability, the unit should be equipped with its automatic exposure timer with 5-second to 60-minute setting at 1-second intervals. The cabinet has adjustable shelf levels for film-to-source distances of 31–61 mm and is fully lead lined to shield the operator from X-rays. A special door interlock safety device automatically turns off the X-ray beam if the door is opened during operation. This is an instrument that uses X-ray film (Kodak X-OMAT 2, Ready Pak; Kodak Ltd) but newer units produce digital and real-time images. When using a Faxitron for decalcification checks, a bone slab should be first radiographed using the automatic exposure timer, and then exposure time, kV, and mA is recorded. This eliminates guesswork for a first exposure and provides the correct exposure time and kV for a repeat radiograph or for subsequent manual exposures of adjacent bone slabs of the same thickness.
Exposure time is dependent on specimen thickness (thicker slabs require longer exposures), film-to-source distance (longer distance requires more time), and type of film used. Large, whole bones, e.g. proximal end of femur with metal prosthesis, can be radiographed using X-OMAT 2, FTSD (upper shelf), longer exposure time (approx. 8 minutes), and higher kV (70 kV). Soft tissues, cartilage, and tumor are more easily seen in underexposed radiographs, useful for evaluating surrounding soft tissue involvement by a bone tumor, e.g. osteosarcoma. The X-ray film can be developed quickly in a radiology department and viewed without delay.
In urgent cases of suspected tumor or infection, an attempt should be made to select a sample with the least mineralization in order to provide the quickest possible diagnosis. These pieces can be fixed, rapidly decalcified, and processed to meet urgent clinical requirements.
The ideal thickness of larger bone pieces is 3–5 mm. If bone slabs are too thick, both decalcification and processing are prolonged while overly thin bone slabs (less than 2 mm) tend to become brittle and bend during processing and embedding which may cause the tissue to pop out of the paraffin block during sectioning. The dense collagen matrix tends to prevent adequate paraffin wax penetration and thin pieces are not held firmly in the softer paraffin embedding media during microtomy.
In order to obtain satisfactory paraffin sections of bone, inorganic calcium must be removed from the organic collagen matrix, calcified cartilage, and surrounding tissues. This is called decalcification and is carried out by chemical agents, either acids to form soluble calcium salts, or chelating agents that bind to calcium ions. Even after decalcification, the dense collagen of cortical bone is remarkably tough and tends to harden more after paraffin processing. Occasionally, small foci of calcifications in paraffin-embedded or frozen tissues can be sectioned without much noticeable damage to the knife or disruption of surrounding tissue. After hematoxylin staining, these foci usually appear cracked and as dark purple granular masses with lighter purple halos.
Any acid, however well buffered, has some damaging effects on tissue stain-aridity. This problem increases with acidity of solutions, i.e. lower pH, and length of decalcification period. Consequently, the rapid decalcifiers are more likely to adversely affect any subsequent staining, especially if not fixed completely. This is most noticeable in cell nuclei with the failure of nuclear chromatin to take up hematoxylin and other basic dyes as readily as soft tissues never exposed to acid solutions. The staining using acid dyes is also less affected, but eosin (an acid dye) can stain tissue a deep, unpleasant, brick red without the preferred three differential shades. These effects on H&E staining can be reduced by doing the decalcification endpoint test, post-decalcification acid removal, and adjustment of the stain procedure.
As noted previously there are two major types of decalcifying agent, i.e. acids and chelating agents, although Gray (1954) lists over 50 different mixtures. Many of these mixtures were developed for special purposes with one used as a fixing and dehydrating agent. Other mixtures contain reagents, e.g. buffer salts, chromic acid, formalin, or ethanol, intended to counteract the undesirable swelling effects that acids have on tissues. Many popular mixtures used today are from the original formulae developed many years ago (Evans & Krajian 1930; Kristensen 1948; Clayden 1952). For most practical purposes, today’s laboratories seem to prefer simpler solutions for routine work. Provided the bone is totally fixed and treated with a decalcifier suitable for removal of the amount of mineral present, the simple mixtures work as well or better than more complex mixtures.
Acid decalcifiers can be divided into two groups: strong (inorganic) and weak (organic) acids. As Brain (1966) suggested, many laboratories keep an acid from each group available for either rapid diagnostic or slower, routine work.
The components in proprietary decalcifying solutions are often trade secrets. Manufacturers provide Material Safety Data Sheets (MSDS) that frequently indicate the type and concentration of acid. Their product data sheets usually indicate if a solution is rapid or slow, and give decalcification instructions and warnings against prolonged use. Rapid proprietary solutions usually contain hydrochloric acid (HCl), whereas slow proprietary mixtures contain buffered formic acid or formalin/formic acid. A study (Callis & Sterchi 1998) found that dilution of a proprietary HCl solution was not deleterious for effective decalcification or staining, and this is an option if a strong mixture is considered too concentrated. Chelating reagents such as EDTA mixtures are also available pre-mixed. Although proprietary mixtures have no obvious advantages over solutions prepared in laboratories, their use is increasingly popular in busy laboratories because they are reliable, time and cost-effective while addressing some safety issues by eliminating handling and storage of concentrated acids.
These may be used as simple aqueous solutions with recommended concentrations of 5–10%. They decalcify rapidly, cause tissue swelling, and can seriously damage tissue stainability if used longer than 24–48 hours. Old nitric acid is particularly damaging, and should be replaced with fresh stock. Strong acids, however, tend to be more damaging to tissue antigens for immunohistochemical staining, and enzymes may be totally lost.
Strong acids are used for needle and small biopsy specimens to permit rapid diagnosis within 24 hours or less. They can be used for large or heavily mineralized cortical bone specimens with decalcification progress carefully monitored by a decalcification endpoint test (Callis & Sterchi 1998). The following is a list of strong acid decalcifying solutions. The formulae and preparations are available in Bancroft & Gamble, sixth ed. (2008).
Of these, formic is the only weak acid used extensively as a primary decalcifier. Acetic and picric acids cause tissue swelling and are not used alone as decalcifiers but are found as components in Carnoy’s, Bouin’s, and Zenker’s fixatives. These fixatives will act as incidental, although weak, decalcifiers and could be used in urgent cases with only minimal calcification. Formic acid solutions can be aqueous (5–10%), buffered or combined with formalin. The formalin–10% formic acid mixture simultaneously fixes and decalcifies, and is recommended for very small bone pieces or needle biopsies. However, it is still advisable to have complete fixation before any acid decalcifier is used. The salts, sodium formate (Kristensen 1948) or sodium citrate (Evans & Krajian 1930), are added to formic acid solutions making ‘acidic’ buffers. Buffering is used to counteract the injurious effects of the acid. However, in addition to low 4–5% formic acid concentration, increased time is needed for complete decalcification. Formic acid is gentler and slower than HCl or nitric acids, and is suitable for most routine surgical specimens, particularly when immunohistochemical staining is needed. Formic acid can still damage tissue, antigens, and enzyme staining, and should be endpoint tested. Decalcification is usually complete in 1–10 days, depending on the size, type of bone, and acid concentration. Dense cortical or large bones have been effectively decalcified with 15% aqueous formic acid and a 4% hydrochloric acid–4% formic acid mixture (Callis & Sterchi 1998). The following is a list of weak acid decalcifying solutions. The formulae and preparations are available in Bancroft & Gamble, sixth ed. (2008).
The chelating agent generally used for decalcification is ethylenediaminetetraacetic acid (EDTA). Although EDTA is nominally ‘acidic’, it does not act like inorganic or organic acids but binds metallic ions, notably calcium and magnesium. EDTA will not bind to calcium below pH 3 and is faster at pH 7–7.4; even though pH 8 and above gives optimal binding, the higher pH may damage alkali-sensitive protein linkages (Callis & Sterchi 1998). EDTA binds to ionized calcium on the outside of the apatite crystal and as this layer becomes depleted more calcium ions reform from within; the crystal becomes progressively smaller during decalcification. This is a very slow process that does not damage tissues or their stainability. When time permits, EDTA is an excellent bone decalcifier for enzyme staining, and electron microscopy. Enzymes require specific pH conditions in order to maintain activity, and EDTA solutions can be adjusted to a specific pH for enzyme staining. EDTA does inactivate alkaline phosphatase but activity can be restored by addition of magnesium chloride.
EDTA and EDTA disodium salt (10%) or EDTA tetrasodium salt (14%) are approaching saturation and can be simple aqueous or buffered solutions at a neutral pH of 7–7.4, or added to formalin. EDTA tetrasodium solution is alkaline, and the pH should be adjusted to 7.4 using concentrated acetic acid. The time required to totally decalcify dense cortical bone may be 6–8 weeks or longer, although small bone spicules may be decalcified in less than a week. Formulae and preparations are available in Bancroft & Gamble, sixth ed. (2008).
Several factors influence the rate of decalcification, and there are ways to speed up or slow down this process. The concentration and volume of the active reagent, including the temperature at which the reaction takes place, are important at all times. Other factors that contribute to how fast bone decalcifies are the age of the patient, type of bone, size of specimen, and solution agitation. Mature cortical bone decalcifies slower than immature, developing cortical or trabecular bone. Another factor of mature bone is that the marrow may contain more adipose cells than a young bone. This requires diligent attention to make sure specimens stay immersed in decalcification solution. Of all the above factors, the effectiveness of agitation is still open to debate.
Generally, more concentrated acid solutions decalcify bone more rapidly but are more harmful to the tissue. This is particularly true of aqueous acid solutions, as various additives, e.g. alcohol or buffers, which protect tissues, may slow down the decalcification rate. Remembering that 1 N and 1 M solutions of HCl, nitric, or formic acid are equivalent, Brain (1966) found that 4 M formic acid decalcified twice as fast as a 1 M solution without harming tissue staining, and felt it was advantageous to use the concentrated formic acid mixture. With combination fixative-acid decalcifying solutions, the decalcification rate cannot exceed the fixation rate or the acid will damage or macerate the tissue before fixation is complete. Consequently, decalcifying mixtures should be compromises that balance the desirable effects (e.g. speed) with the undesirable effects (e.g. maceration, impaired staining).
In all cases, total depletion of an acid or chelator by their reaction with calcium must be avoided. This is accomplished by using a large volume of fluid compared with the volume of tissue (20 : 1 is recommended), and by changing the fluid several times during the decalcification process. Brain, however, pointed out that if a sufficiently large volume of fluid is used (100 ml per g of tissue) it is not necessary to renew the decalcifying agent because depletion is less apparent in a larger volume. Small number and similar-sized specimens in one container is preferred.
Ideally, acid solutions should be endpoint tested and changed daily to ensure the decalcifying agent is renewed and that tissues are not left in acids too long or overexposed to acids (i.e. ‘over-decalcification’).
Increased temperature accelerates many chemical reactions, including decalcification, but it also increases the damaging effects which acids have on tissue so that, at 60°C, the bone, soft tissues, and cells may become completely macerated almost as soon as they are decalcified.
The optimal temperature for acid decalcification has not been determined, although Smith (1962) suggested 25°C as the standard temperature, but in practice a room temperature (RT) range of 18–30°C is acceptable. Conversely, lower temperature decreases reaction rates and Wallington (1972) suggested that tissues not completely decalcified at the end of a working week could be left in acid at 4°C over a weekend. This practice may result in ‘over-decalcification’ of tissues, even with formic acid. A better recommendation is to interrupt decalcification by briefly rinsing acid off bone, immersing it in NBF, removing from fixative, rinsing off the fixative, and resuming decalcification on the next working day. Microwave, sonication, and electrolytic methods produce heat, and must be carefully monitored to prevent excessive temperatures that damage tissue (Callis & Sterchi 1998).
Increased temperature also accelerates EDTA decalcification without the risk of maceration. However, it may not be acceptable for preservation of heat-sensitive antigens, enzymes, or electron microscopy work. Brain (1966) saw no objection to decalcifying with EDTA at 60°C if the bone was well fixed.
The effect of agitation on decalcification remains controversial even though it is generally accepted that mechanical agitation influences fluid exchange within as well as around tissues with other reagents. Therefore, it would be a logical assumption that agitation speeds up decalcification, and studies have been done which attempt to confirm this theory. Russell (1963) used a tissue processor motor rotating at one revolution per minute and reported the decalcification period was reduced from 5 days to 1 day. Others, including Clayden (1952), Brain (1966), and Drury and Wallington (1980), repeated or performed similar experiments and failed to find any time reduction. The sonication method vigorously agitates both specimen and fluid, and one study noted cellular debris found on the floor of a container after sonication could possibly be important tissue shaken from the specimen (Callis & Sterchi 1998). Gentle fluid agitation is achieved by low-speed rotation, rocking, stirring, or bubbling air into the solution. Even though findings from various studies are unresolved, agitation is a matter of preference and not harmful as long as tissue components remain intact.
The decalcifying fluid should be able to make contact with all surfaces of a specimen and flat bone slabs should not touch each other or the bottom of a container, as this is enough to prevent good fluid access between the flat surfaces. Bone samples can be separated and suspended in the fluid with a thread or placed inside cloth bags tied with thread or preferably in a cassette. The cassette will provide identification without having to prepare tags for bag suspension. Some workers have cleverly devised perforated plastic platforms to raise samples above a container bottom to permit fluid access to samples.
Ideally, bone should be taken from the acid solution as soon as all calcium has been removed from it. It is still possible that the outer parts of a sample will be overexposed to the acid, although these parts usually stain no differently from inner portions, which are the last to be decalcified. Tissues decalcified in acids for long time periods or in high acid concentrations are more likely to show the effects of over-decalcification, whether or not all mineral has been removed.
Consequently, it is important for a laboratory to control a decalcification procedure by using a decalcification endpoint test to know when calcium removal is complete and, if incomplete, renew the decalcifying agent. If laboratories do not perform endpoint testing, it is recommended they should do so. When using formic, HCl or nitric acids, daily testing is recommended unless near the endpoint; then test every 3–5 hours when possible. With EDTA, weekly tests are sufficient unless solution changes are more frequent. It is recommended that EDTA be changed often at first, since the reaction with calcium is initially quicker and then slows down as the calcium in the tissue is depleted. Minimally calcified tissues and needle biopsies decalcified by a strong acid may be tested only once. It is wise practice for a laboratory that performs a high number of bone biopsy specimens to establish a time range (that depends on the amount of cortical bone present) for ideal decalcification time for bone biopsies. Once the process is established it would eliminate guessing, testing, and handling of those delicate samples. Biopsy decalcification is fairly consistent on multiple-sized biopsies within an hour or two of acid exposure. Wrapping a bone needle biopsy in tissue paper and leaving it wrapped until embedding prevents any loss of cells or tissue during the washing or decalcification process. Sponges sometimes pull cells from a sample when washing and decalcifying. Often these may be urgent cases where a shorter decalcification time is allowed, but the sample must be carefully treated and incomplete decalcification is still possible. If tissue is still slightly under-decalcified after paraffin embedment and sectioning, surface decalcification can be done. Problem blocks should be identified as such so that proper treatment is given should further microtomy be requested.
There are several methods for testing the completion of decalcification, with two considered to be the most reliable. These are specimen radiography, using an X-ray unit and the chemical method to test acids and EDTA solutions. Another method first used to test nitric acid is a weight loss, weight gain procedure that provides relatively good, quick results with all acids and EDTA (Mawhinney et al. 1984; Sanderson et al. 1995). Although still used, ‘physical’ tests are considered inaccurate and damaging to tissues. Probing, ‘needling’, slicing, bending, or squeezing tissue can create artifacts, e.g. needle tracks, disrupt soft tumor from bone, or cause false-positive microfractures of fine trabeculae, a potential misdiagnosis. The ‘bubble’ test is subjective and dependent on worker interpretation.
Methods for chemical testing of acid decalcifying fluids detect the presence of calcium released from bone. When no calcium is found or the result is negative, decalcification is said to be complete and may entail using one extra change of decalcifier after actual completion. EDTA can be chemically endpoint tested by acidifying the used solution; this forces EDTA to release calcium for precipitation by ammonium oxalate (Rosen 1981).
Calcium oxalate test (Clayden 1952)
This method involves the detection of calcium in acid solutions by precipitation of insoluble calcium hydroxide or calcium oxalate, but is unsuitable for solutions containing over 10% acid even though these could be diluted and result in a less sensitive test.
If a white precipitate (calcium hydroxide) forms immediately after adding the ammonium hydroxide, a large quantity of calcium is present, making it unnecessary to proceed further to step 3 which would also be positive. Testing can be stopped and a change to fresh decalcifying solution made at this point. If step 2 is negative or clear after adding ammonium hydroxide, then proceed to step 3 to add ammonium oxalate. If precipitation occurs after adding the ammonium oxalate, less calcium is present. When a smaller amount of calcium is present, it takes longer to form a precipitate in the fluid, so, if the fluid remains clear after 30 minutes, it is safe to assume decalcification is complete.
‘Bubble’ test. Acids react with calcium carbonate in bone to produce carbon dioxide, seen as a layer of bubbles on the bone surface. The bubbles disperse with agitation or shaking but reform, becoming smaller as less calcium carbonate is reduced. As an endpoint test, a bubble test is subjective and unreliable, but can be used as a guide to check the progress of decalcification, i.e. tiny bubbles indicate less calcium present.
Radiography. This is the most sensitive test for detecting calcium in bone or tissue calcification. The method is the same as specimen radiography using a FAXITRON with a manual exposure setting of approximately 1 minute, 30 kV, and Kodak X-OMAT X-ray film on the bottom shelf. It is possible to expose several specimens at the same time. The method is to rinse acid from the sample, carefully place identified bones on waterproof polyethylene sheet on top of the X-ray film, expose according to directions, and leave bones in place until the film is developed and examined for calcifications. Bones with irregular shapes and variable thickness can occasionally mislead workers on interpretation of results. This problem is resolved by comparing the test radiograph to the pre-decalcification specimen radiograph and correlate suspected calcified areas with specimen variations. Areas of mineralization are easily identified, although tiny calcifications are best viewed using a hand-held magnifier. Metal dust particles from saw blades are radio-opaque, sharply delineated fragments that never change in size. These are unaffected by decalcification, appearing as gray specks on the bone surface and can be easily removed. Spicules of metal, metallic paint, or glass forced deep into tissue by a traumatic injury are also sharply delineated but cannot be removed without damaging the tissue. Radiography only indicates the presence of deeper foreign objects, and care must be taken during microtomy to not damage the knife (Fig. 16.5).
Acids can be removed from tissues or neutralized chemically after decalcification is complete. Chemical neutralization is accomplished by immersing decalcified bone into either saturated lithium carbonate solution or 5–10% aqueous sodium bicarbonate solution for several hours. Many laboratories simply rinse the specimens with running tap water for a period of time. Culling (1974) recommended washing in two changes of 70% alcohol for 12–18 hours before continuing with dehydration in processing, a way to avoid contamination of dehydration solvents even though the dehydration process would remove the acid along with the water.
Adequate water rinsing can generally be done in 30 minutes for small samples and larger bones in 1–4 hours in order to not delay processing. Samples needing immediate processing, e.g. needle biopsies, can be blotted or quickly rinsed to remove acid from surfaces before proceeding to the first dehydrating fluid. It is important to avoid contaminating the first dehydrating fluid with acids, and washing bones even for a short time is good practice particularly with large bone slabs.
Acid-decalcified tissues for cryomicrotomy must be thoroughly washed in water or stored in formal saline containing minimal amounts of sucrose (3–10%), or PBS with 3–10% sucrose, at 4°C before freezing. This helps avoid any residual acid in the tissue from corroding the metal knife. Caution: higher percentages of sucrose may prevent the tissue from fully freezing or freezing unevenly, causing soft spots in the tissue which will create thick-thin appearing sections or the tissue will fall out due to improper freezing.
Tissues decalcified in EDTA solutions should not be placed directly into 70% alcohol, as this causes residual EDTA to precipitate in the alcohol and within the tissue. The precipitate does not appear to affect tissue staining since EDTA is washed out during these procedures, but may be noticeable during microtomy or storage when a crystalline crust forms on the block surface. A water rinse after decalcification or overnight storage in formal saline, NBF, or PBS should prevent this.