Aiza Sanmiguel

A 1 year old bearded dragon developed chronic crusty lesions over the body, face and tail. Skin biopsies were taken and submitted to VPG Histology for examination. This revealed epidermal hyperplasia and prominent hyperkeratosis, areas of superficial epidermal necrosis and a mild inflammatory infiltrate within the epidermis and dermis. Faint outlines of structures suspected to represent fungal hyphae and spores were observed in the keratin layer.

The skin biopsy showing hyperplasia of the epidermis (E) and a prominent keratin layer (K).

A PAS stain revealed numerous fungal hyphae (long arrows) and conidia (short arrows) in the keratin layer.

These findings indicate a fungal infection. The histological features of the fungal organisms, together with the clinical presentation support a diagnosis of “yellow fungus disease”.

‘Yellow fungus disease’ is a dermatomycosis of bearded dragons. The disease presents initially with crusty lesions of hyperkeratosis, progressing to yellow or brown discoloured areas and eventually cutaneous necrosis. Areas of necrosis may slough, resulting in ulcerated lesions. Progression to underlying tissues and systemic dissemination may occur (1).

A fungus previously termed ‘Chrysosporium anamorph of Nannizziopsis vriesii’ (CANV) has been isolated from such lesions (2) . Fungi with similar features have also been identified in other reptile species, including green iguanas, chameleons, snakes and crocodiles. Further analysis of these isolates has led to reclassification of the fungi, and the species most commonly associated with yellow fungus disease in bearded dragons is now considered to be Nannizziopsis guarroi (3,4). Other species previously classified as CANV include Nannizziopsis dermatitidis isolated from chameleons, N. crocodilii (saltwater crocodiles), Paranannizziopsis australasiensis (bearded dragons and snakes) and Ophidiomyces opiodiicola (snakes).

Unlike many other fungi, which act as opportunistic pathogens, fungi of the former CANV-complex are considered primary pathogens of reptiles and are contagious. Stress and overcrowding are predisposing factors.

A minimum of 2 minutes of exposure to a 10% dilution of commercial bleach has been recommended for disinfection of surfaces and instruments contaminated with N. guarroi (5)

Histopathological examination of skin biopsies identifies fungal hyphae and conidia. Further confirmation can be obtained by using fungal culture or PCR analysis.

The zoonotic risk is considered to be very low. Rare cases of infections with fungi, suspected to represent a CANV-like species have been reported in immunocompromised patients, but further analysis indicated that these may represent a distinct lineage, separate from the reptile species. Nevertheless, reptiles may be carriers of a range of pathogens, including Salmonella spp.,  and good hygiene is therefore essential in particular in households with immunocompromised individuals.

1. Pare J., Sigler L. Fungal Diseases in Mader’s Reptile and Amphibian Medicine and Surgery.Saunders Elsevier; St. Louis, Missouri, USA: 2006. pp. 217–226.
2. Bowman MR, Paré JA, Sigler L, Naeser JP, Sladky KK, Hanley CS, Helmer P, Phillips LA, Brower A, Porter R. (2007).Deep fungal dermatitis in three inland bearded dragons (Pogona vitticeps) caused by the Chrysosporium anamorph of Nannizziopsis vriesii. Med Mycol. Jun;45(4):371-6.
3. Schilliger L, Paillusseau C, François C, Bonwitt J. (2023) Major Emerging Fungal Diseases of Reptiles and Amphibians. Pathogens. Mar 8;12(3):429.
4. Sigler L, Hambleton S, Paré JA. (2013). Molecular characterization of reptile pathogens currently known as members of the chrysosporium anamorph of Nannizziopsis vriesii complex and relationship with some human-associated isolates. J Clin Microbiol. 51(10):3338-57.
5. Jourdan B., Hemby C., Allender M.C., Levy I., Foltin E., Keller K.A. (2023) Effectiveness of Common Disinfecting Agents Against Isolates of Nannizziopsis guarroi.  Herpetol. Med. Surg. 33(1), 40-44.

Leo, 1 year old male Domestic Shorthair cat

Clinical history

Leo presented with swelling of the left forepaw characterised by the presence of multiple cutaneous lesions with scabs and poor skin integrity.

Histology

Three punch biopsies of haired skin were submitted for histopathological examination. These samples showed extensive epidermal ulceration, and severe epidermal and dermal necrosis, characterized by the presence of large amounts of amorphous eosinophilic material, pyknotic and karyorrhectic cellular debris, degenerate and viable neutrophils, fibrin and haemorrhage. There was marked inflammation extending from the ulcerated tissue into the deep dermis comprising large numbers of neutrophils and eosinophils, moderate to large numbers of macrophages often palisading around necrotic and disrupted hair follicles which faded in the face of necrosis and the associated inflammatory infiltrates (Figure 1). Fragments of free hair shafts were frequently entrapped within the inflamed tissue. There was also mild to moderate oedema and proliferation of endothelial cells and fibroblasts in the background.

In the superficial disrupted tissue there were multiple residual islands of follicular infundibular epithelium embedded within the areas of necrosis. Keratinocytes often showed variably-sized bright eosinophilic intracytoplasmic inclusions, consistent with cowpox A-type inclusion bodies (Figure 2).

Photomicrograph showing severe ulcerative and necrotising dermatitis (HE, 10x).

Figure 2: Photomicrograph showing large numbers of variably-sized bright eosinophilic intracytoplasmic viral inclusion bodies (arrows) within keratinocytes, consistent with cowpox A-type inclusion bodies (HE, 40x).

Interpretation

Dermatitis, ulcerative, necrotising, neutrophilic, eosinophilic, histiocytic to pyogranulomatous, multifocal, subacute, severe, with intralesional eosinophilic intracytoplasmic inclusion bodies in keratinocytes – consistent with cowpox A-type inclusion bodies.

Molecular testing

A fresh skin biopsy was also submitted to our lab for Orthopoxvirus qPCR, and yielded a positive result, with a Ct value of 15.5. Ct values typically range from 15-40, and the higher the Ct value, the lower the amount of DNA present in the sample. Therefore, these results indicate a high level of Orthopoxvirus DNA in the sample submitted.

 Comments

Histology of the samples of haired skin submitted captured a severe ulcerative and necrotising dermatitis (Figure 1). Residual hair follicles were observed, and keratinocytes showed moderate to large numbers of variably-sized and bright eosinophilic intracytoplasmic inclusion bodies – consistent with cowpox A-type inclusion bodies (Figure 2). In this case, cowpox virus infection was presumptively diagnosed based on the presence of characteristic intracytoplasmic inclusion bodies, and was confirmed using Orthopoxvirus qPCR.

Ulcerative and necrotizing dermatitis in cats can also be attributed to feline herpesviral dermatitis, resulting from Felid herpesvirus 1 infection. It’s essential to consider this as a potential alternative diagnosis, particularly in cases where eosinophilic intracytoplasmic inclusion bodies are absent, and if the mixed dermal inflammatory infiltrate includes eosinophils.

What is cowpox?

Cowpox is an infection caused by cowpox virus (genus Orthopoxvirus), considered a re-emerging zoonotic pathogen and a public health threat due to increasing numbers of cases in humans and animals in Europe over the past decade, including within the United Kingdom. In addition to cattle, cowpox virus may infect cats, dogs, and various non-domestic species. Serological studies have established that the most likely reservoirs of CPXV in Great Britain are bank voles (Myodes glareolus), short-tailed field voles (Microtus agrestis) and, to a lesser extent, wood mice (Apodemus sylvaticus). The highest seasonal incidence of feline cowpox is from late summer to autumn, corresponding with the peak size of rodent populations.

The majority of feline cowpox cases feature a solitary primary cutaneous lesion, typically found on the head, neck, forelimb, or paws. During the viraemic phase, which occurs 1-3 weeks after the appearance of the primary lesion, secondary skin lesions emerge widely. Initially, the skin lesions present as small papules, which gradually enlarge into nodules and subsequently ulcerate, forming craters and crusts. Vesicles are seldom observed, particularly on the oral mucosa and inner pinna. Generally, full recovery is observed within 4-5 weeks.

Atypical cutaneous cowpox virus infection has also been observed in cats, presenting with skin lesions on the distal limbs characterised primarily by oedema, hyperaemia and plaque-like lesions, rather than the typical ulcerated papules. Feline cowpox can also manifest in a mucocutaneous form and, in rare instances, can become systemic, replicating in internal organs, particularly the lungs and upper gastrointestinal tract, often resulting in a fatal outcome. Rare cases of fatal necrotizing pneumonia without the presence of typical skin lesions have also been reported.

Cowpox has rarely been reported in dogs and other domestic species. However, non-domestic species are also susceptible, with exotic felids, particularly cheetahs, being at high risk of developing systemic disease.

Zoonotic potential

Cowpox is a rare zoonotic disease that poses a significant risk for immunocompromised individuals, including children, elderly and pregnant women. Approximately 50% of human cowpox cases are associated with feline cowpox infections.¹ Therefore, pet owners and veterinary staff who come into direct contact with infected cats are advised to seek medical attention if they develop skin lesions suggestive of cowpox.

References

¹Lawn, R. (2010) Risk of cowpox to small animal practitioners. Veterinary Record, 166: 631-631. https://doi.org/10.1136/vr.c2505

Luna, a 6-year-old cross breed female entire dog

Clinical history

A typical clinical history for a CRGV case includes development of one or multiple areas of skin ulceration, often on the abdomen and limbs, followed by lethargy, anorexia, renal injury and loss of function. Samples from the skin and kidney are often submitted post-mortem.

Histology

Histologically the epidermis presents with areas of ulceration and necrosis. Fibrinoid degeneration and vasculitis of the dermal and subcutaneous vessel are often observed. Thrombi formation is also a frequent finding.

Figure 1: Epidermal ulceration and necrosis (green arrow) with overlying serocellular crust (black arrow) with fibrinoid vasculitis of dermal vessels (black circles).

Within the sections of the kidney there is usually extensive glomerular damage. Glomeruli often are hypereosinophilic with loss of cellular details and thickening and distortion of capillary loops and basement membrane with peripheral adhesion to the Bowman’s capsule. Glomerular sclerosis can be observed. Tubules are multifocally lined by degenerate epithelial cells and protein casts are multifocally observed within the lumen. Fibrinoid degeneration of blood vessels is also observed.

Figure 2: Hypereosinophilic glomeruli with diffusely thickened basement membranes and adhesions to the Bowman’s capsule (black circle) and sclerotic glomeruli (more advanced granular degeneration) (green circle).

Figure 3: Sclerotic glomeruli (green circle), tubule lined by degenerate epithelium (green arrow) and tubules containing protein material within the lumen (black arrow).

Cutaneous and renal glomerular vasculopathy (CRGV) is a rare condition that affects dogs only, causing skin lesions and severe kidney damage. This disease is also known as “Alabama Rot,” and it was first identified in Greyhounds in the United States in the 1980s but has since been reported in various breeds, but Hounds, gundogs and pastoral dogs are often overrepresented. CRGV is recognised mainly in the UK, however, a few isolated cases have been reported in Germany and the Republic of Ireland. This disease is characterised by seasonal outbreaks with most cases reported between the months of November and May. Woodland habitat, increased winter temperatures, as well as high rain falls, have been recognised as important factors in the distribution of this disease. Some dogs with CRGV are reported to have been in touch with another dog with it and often multiple dogs in the same household can be affected.

Despite extensive research, the exact cause of CRGV remains elusive and the condition often presents a diagnostic and therapeutic challenge for veterinarians in the UK.

CRGV typically begins with skin lesions, particularly on the limbs, abdomen, and muzzle and these patients are often lame. Skin lesions usually consist of ulcers or erosions, accompanied by oedema, and pain. Affected dogs may also exhibit signs of systemic illness, such as lethargy, anorexia, vomiting, and fever. As the disease progresses, renal injury and eventually renal failure occur.

The pathogenesis of CRGV is not fully understood, but it involves damage to small blood vessels, particularly the glomerular capillaries in the kidneys, leading to endothelial damage, fibrin deposition, thrombi formation, ischemia and organ damage with subsequent loss of function. Currently there is no single reliable test for an ante-mortem diagnosis of CRGV in dogs and histopathology examination of the kidney remains the gold standard for identification of the thrombotic microangiopathy, which is considered the histopathological hallmark of this disease. For this reason, samples are often submitted post-mortem for confirmation of a clinical suspicion.

Ingestion of bacteria-associated Shiga toxins has been proposed as a possible underlying cause, however, studies have failed to consistently identify the presence of toxins, bacteria, viruses or other microorganisms in the affected tissues.

Diagnosis of CRGV relies on a combination of typical clinical signs, biochemical tests, and histopathological evaluation. Skin biopsies can be helpful and reveal changes consistent with vasculopathy, which may help the clinician identify this disease.

References:

Walker JJA, Holm LP, Sarmiento ÓG, Caianiello R, Cortellini S, Walker DJ. Clinicopathological features of cutaneous and renal glomerular vasculopathy in 178 dogs. Vet Rec. 2021 Aug;189(4):e72. doi: 10.1002/vetr.72. Epub 2021 Jan 24. PMID: 33829498.

Holm LP, Stevens KB, Walker DJ. Pathology and Epidemiology of Cutaneous and Renal Glomerular Vasculopathy in Dogs. J Comp Pathol. 2020 Apr;176:156-161. doi: 10.1016/j.jcpa.2020.03.003. Epub 2020 Apr 8. PMID: 32359630.

Hope A, Martinez C, Cassidy JP, Gallagher B, Mooney CT. Canine cutaneous and renal glomerular vasculopathy in the Republic of Ireland: a description of three cases. Ir Vet J. 2019 Nov 16;72:13. doi: 10.1186/s13620-019-0151-7. PMID: 31762988; PMCID: PMC6858974.

Our CEO, Fiona Gosling, has co-authored an article titled “Clinicopathological and Diagnostic Imaging Findings in a Dog with Neurocandidiasis,”

The article was published in the Journal of Veterinary Internal Medicine on 20th May 2024.

Read the full article here

Ben, a 2 year old Marmoset monkey.

Clinical history

Ben had a history of a fluid-filled swelling in the mammary area. Previously, fluid was sampled and was found to be of low cellularity. Lesions progressed with erythematous skin and secondary pyoderma over the entire ventrum and thickened, sore scrotal skin.

Histology

The biopsies submitted were representative of subcutaneous adipose and some fibrous connective tissue. Dissecting between the adipose and fibrous connective tissue were variably ectatic irregular and branching channels filled with clear space and rarely some erythrocytes. These channels were lined by flat and histologically unremarkable endothelial cells. No mitotic figures were observed in 2.37 mm2 that were examined (equivalent to 10 high power fields (x400)).

Fig. 1: The biopsy is composed of subcutaneous adipose and fibrous connective tissue. Empty channels dissecting between the tissue are already observed at low magnification (x10); examples of channels marked with arrowheads.

Fig. 2: Higher magnification of channels (x200). The channels are mostly empty, only rare erythrocytes are observed (blue arrowhead). The channels are lined by histologically unremarkable endothelial cells (red arrowhead).

Interpretation

Consistent with lymphangiomatosis, skin, mammary area.

Note: Definitive confirmation of a lymphatic proliferation requires IHC for the lymphatic endothelial marker LYVE-1. However, based on the combination of clinical history and histologic appearance, lymphangiomatosis was considered the most likely diagnosis.

What is Lymphangiomatosis?

Lymphangiomatosis is a rare congenital disorder thought to represent a malformation arising from failure of primitive lymphatic systems to adequately separate from or communicate with the venous system. Though the disorder is not thought to be neoplastic, progression and recurrence frequently occur.

In humans the disorder occurs primarily in children and rarely manifests after the first two decades.

In animals, lymphatic malformations are very rare. The reports available mostly describe cutaneous lymphangiomatosis in dogs and cats with rare reports in other species. The lesions are most frequently observed on the abdomen, ventral neck, inguinal area, prepuce and legs. Other lymphatic malformations are rarely reported in dogs and have been observed in the retroperitoneum, abdominal cavity, spleen and liver. As in humans, mostly young animals are affected, although a case of lymphangiomatosis in an old dog has been published.

Cutaneous lymphangiomatosis clinically presents as poorly demarcated, fluctuant swellings, that are often progressive. Tracts and ulcers may occur and may drain serous fluid.

Histologically, the lesion is characterised by a poorly circumscribed mass composed of angular, dilated and partially interconnected channels, which may contain some proteinaceous fluid; erythrocytes are rarely observed. The channels are lined by inconspicuous endothelium with no cellular atypia and minimal or no mitotic activity.

Lymphangiomatosis must be differentiated from clinically and histologically well-demarcated lymphangiomas, for which progression and recurrence after complete local resection would not be expected.

To the best of the author’s knowledge, lymphangiomatosis has not been described in marmosets, so far.

References

Gross et al.: Skin diseases of the dog and cat. Clinical and histopathological diagnosis. 2^nd edition (2005)

Ask JPC. Conference 24 – 2010: case 0120110316

Park et al.: Case report: Generalized lymphatic anomaly of multiple abdominal organs in a young dog. Front Vet Sci (2023)

Berry et al.: Lymphangiomatosis of the pelvic limb in a Maltese dog. J Small Animal Pract. (1996)

Maeda S, Fujino Y, Tamamoto C, Suzuki S, Fujita A, Takahashi M, et al. Lymphangiomatosis of the systemic skin in an old dog. J Vet Med Sci. (2013)

Driessen F, Cushing T, Baines SJ. Retroperitoneal lymphatic malformation in a dog. Acta Vet Scand. (2020)

Locker SH, Maxwell EA, Vilaplana Grosso F, Bertran J, Shiomitsu K. Novel treatment of recurrent abdominal lymphatic malformations in a dog. Vet Rec Case Rep. (2021)

Belanger MC, Mikaelian I, Girard C, Daminet S. Invasive multiple lymphangiomas in a young dog. J Am Anim Hosp Assoc. (1999)

Ramirez GA, Sanchez-Salguero X, Molin J. Primary Cystic Lymphangioma of the spleen in an adult dog. J Comp Pathol. (2020)

For surgery of a neoplasm with curative intent, the goal is to achieve a macroscopic margin around the tumour at surgery and a microscopically tumour-free margin. The assumption is that with an ‘adequate’ margin, local recurrence can be prevented. In reality, the matter is more complex. Neoplasms with neoplastic tissue histologically extending to the tissue border (often referred to as ‘dirty’ margin) may not recur, while in other cases neoplasms with microscopic tumour-free margins (often referred to as ‘clean’ margins) may locally recur. Possible reasons for this are discussed in the paragraphs below.

“Dirty” margins: Why hasn’t the tumour recurred?

The reason for non-recurrence of neoplasms with ‘dirty’ histologic margins may be a false positive result due to a tissue artefact (more explanations on artefacts below).

It could also be that a tumour was ‘shelled-out’. Histologically, there may be no non-neoplastic tissue visible surrounding the neoplasm, however, this does not mean that tumour cells have remained in situ. Even if some neoplastic tissue remains at the surgical site, it may be eradicated by the host immune response (possibly triggered by the surgical procedure).

For benign, well-circumscribed neoplasms, ‘shelling-out’ is generally expected to be curative and literature indicates, that for some ‘low grade’ malignancies, such as ‘low grade’ soft tissue sarcoma, in many cases local recurrence does not occur after ‘shelling-out’.

For some low grade malignancies with incomplete local resection, local recurrence may be delayed and may still occur after a longer observation period.

“Clean” margins: Why has the tumour recurred?

Only a small subsection of the tumour and its marginal tissue are assessed histologically. Therefore, while the margins may be “clean” in the examined sections, there may be neoplastic tissue at the surgeon-cut edge in areas not histologically assessed. This is particularly problematic in highly infiltrative neoplasms.

Some neoplasm (e.g. high grade canine cutaneous mast cell tumours) tend to form satellite tumours (local metastasis) in the vicinity, which may not be resected with the primary tumour.

Theoretically, there may also be a residual pro-tumorigenic microenvironment at the excision site. This could predispose to novel malignant transformation at the site or induce “homing” signals that draw circulating neoplastic cells to the tumour environment.

Interpretation of the Margin Assessment

The histologic assessment of the margins must be interpreted in conjunction with clinical and surgical features, including location of the mass, the surgical margin and demarcation at surgery. Additionally, the specific tumour biology is important with some types of neoplasms having higher potential for recurrence regardless of the histologic tumour free distance. Therefore, correlation with tumour type and grade is critical. This information, along with other clinical factors, will need to be integrated to formulate the treatment plan, including the advisability of revision surgery, radiotherapy, and chemotherapy.

Outlined below is information on the terminology and techniques that we use in our lab for histologic margin assessment. Additionally, there are limitations of histologic margin assessment of which pathologists and clinicians should be aware.

Terminology

Histologic margin: The tissue that separates the neoplastic tissue from the tissue border. Often, the tissue border is referred to as the margin (as in the neoplasm extends to the margin); however, this is a wrong use of terminology. Please be aware that we only refer to the tissue border as the ‘surgeon-cut edge’ if the tissue was inked by the surgeon. This is because we cannot know if the tissue border we ink in the lab is the actual surgeon-cut edge (more explanations see below).

Histologically tumour-free distance: the width of the margin.

Fig. 1: Cross section through a mast cell tumour; deep tissue border inked green by lab.

Margin dimensions: the deep margin is the tissue underlying the mass. We refer to the margins to the sides of the mass as horizontal. You may also see these margins referred to as lateral. We prefer to use the term “horizontal” because lateral is also a descriptor that may more precisely indicate location (as in lateral and medial). If the sample can be orientated and a more specific location can be indicated for the horizontal margins, we use the usual terminology (cranial/ rostral, caudal, ventral, dorsal, lateral, medial, etc.).

NB: Using sutures is a helpful method to provide orientation. In order to provide the most accurate assessment of orientation, please indicate the orientation in at least two different planes. For example, for a mass on the left lateral thorax, place 1 suture to indicate the dorsal horizontal border and 2 sutures to indicate the cranial tissue border. We will be able to provide you with a more precise assessment of the histologic tumour free distance at specific parts of the sample with this information.

Cruciate sections: A sample is sectioned cross-wise resulting in a transverse and longitudinal plane of section (see also below in techniques).

Shaved/ ‘en-face’ margins: The peripheral tissue of a sample is shaved off and embedded with the surface down in the block to be cut first (see also below in techniques).

Techniques for histologic margin assessment

Cruciate sections

This is the technique most commonly used, especially for ellipsoid excisions of skin tumours. A section is cut transversly through the narrowest area of the sample and then longitudinal sections are cut perpendicularly through the wider ends of the sample. This allows for examination of one plane of section through each of the 4 horizontal dimensions of the sample and examination of the deep margin.

Fig. 2: Illustration of technique to take cruciate sections.

Serial slicing and serial transverse section

For some samples, serial transverse slicing is the more appropriate technique to apply. Examples of samples that are generally serially sliced are small ellipsoid excisions, mammary strips or scar revisions. Depending on the size of the sample all (small ellipsoid excisions) tissue is subsequently processed or representative transverse sections are processed.

Fig. 3: Illustration of serial slicing a sample.

Shaved/ ‘en-face’ margins

As the cruciate and serial transverse sections allow for the examination of only a small proportion of the surface area of the tissue border, shaving of the deep and horizontal tissue border may be applied as an additional method and at an extra cost (HIMASK) charge code. This is particularly useful for neoplasms that appear poorly demarcated/ where infiltrative growth is suspected.

We do not shave neoplasms that have been narrowly excised (‘marginal excision’, ‘shelled-out’) because inevitably, neoplastic tissue will be observed in the shaved sections and no additional useful information is added. If shaving was requested, but is not thought to be useful or possible, we will inform you and remove the charge for shaving.

Fig. 4: Illustration of combination of cruciate sections and shaving tissue borders.

Techniques for complex samples

Complex sample types (such as whole extremities, maxillectomies, mandibulectomies, enterectomies, liver or lung lobes, digits, etc.) mostly require more bespoke methods to allow for histologic margin assessment. This is often a combination of representative sections through the margin into the mass and shaving of tissue borders. This is included in our charge for complex samples (HBEET).

Inking of samples

Inking of tissue borders of a sample can be very helpful for tissue orientation, as well as ensuring that the tissue border on a histologic slide represents the surgeon-cut edge created at surgery.

Inking is best performed by the surgeon. This will help the laboratory technicians and pathologists orientate in particular complex samples. Importantly, any artefact that may be introduced to the margin tissue between surgery and macroscopic examination in the lab can be spotted much easier. Lastly, ink sticks better to fresh tissue than formalin fixed tissue and does not ‘bleed’ into the tissue as much.

Any samples where margin assessment is required that do not come to the laboratory pre-inked are inked in the lab; however, the caveats outlined above need to be kept in mind.

Figure 5A: Green ink applied to tissue border by lab; ink bled into tissue.

Figure 5B: Black ink applied to surgeon-cut edge by surgeon; sharp demarcation, no bleeding into tissue.

Limitations

We can only report the absence of neoplastic cells/ tissue at the tissue border in the planes of tissue that we examine. The assumption is that for most benign tumours and well-demarcated malignant tumours, the absence of neoplastic cells/ tissue at the tissue border correlates well with the completeness of local excision.

For infiltrative neoplasms (e.g. feline injection site sarcoma) and neoplasms that tend to form satellite tumours (e.g. high grade canine cutaneous mast cell tumours), the relationship between the absence of tumour tissue at the tissue border and completeness of local excision is unknown.

Figure 6: Satellite tumour extends to the tissue borders with likely incomplete local excision; neoplastic cells do not extend to the tissue borders in the examined tissue section.

We cannot know if neoplastic cells have remained in situ, even if neoplastic cells are observed at the tissue border histologically. For neoplasms that have been excised with no surrounding non-neoplastic margin (‘shelled out’, ‘marginal excision’)) it is impossible to say if neoplastic cells have been left behind or not. We assume that there is a higher likelihood that tumour cells remain in situ if tumour tissue is observed at the tissue border in tumours that are less demarcated and/or infiltrative.

Figure 7: Well-demarcated mass, neoplastic tissue at tissue border in transverse section (1).

Figure 8: Poorly demarcated, infiltrative mass, neoplastic tissue at tissue border in transverse section (1).

Artefacts

As for all diagnostic tests, we must be aware of artefacts that may occur during processing, which may cause false interpretation of data. Knowing that these artefacts can occur allows us to take pre-emptive measures to reduce their clinical impact.

Tissue shrinkage: This is an expected artefact that changes the extent of the surgical margin compared to the histologically tumour free margin. Most of the shrinkage occurs immediately after surgery due to contraction of the tissue. Some tissue shrinkage is caused by formalin-fixation. The extent of the tissue shrinkage is unpredictable. Tissues may shrink between 10-50% and tissue components may be differently affected (e.g. skeletal muscle and fibrous connective tissue would be expected to shrink more than adipose tissue). This tissue shrinkage may also cause distortion of the sample, which can change the original extent of the margins. This can somewhat (but not completely) be prevented by suturing together the skin and the deep fascial plane (if one could be taken).

Distortion of samples can also be caused by using unsuitable containers. This is most apparent if samples get squeezed into containers that are too small. In addition to the distortion of the tissue borders, placing samples in small containers affects the speed of fixation. Poor fixation can cause other tissue artefacts.

Transport induced: Adipose tissue is very prone to laceration, which can happen during transport of the sample. Unless the surgeon-cut edge has been immediately inked after surgery, it is nearly impossible to differentiate the ‘true’ from a ‘false’ surgeon-cut edge in the lab.

All of these factors may lead to reporting of unexpectedly narrower margins or in the worst case false positive reporting of neoplastic tissue at the tissue border.

If you have any further questions or would like us to arrange a CPD on histologic margin assessment for your team, please complete this short form or contact your nearest laboratory.

References

Bray et al.: Evaluating the relevance of surgical margins. Part one: The problems with current methodology. Vet Comp Oncol 21 (2023).

Bray et al.: Evaluating the relevance of surgical margins. Part two: Strategies to improve prediction of recurrence risk. Vet Comp Oncol 21 (2023)

Milovancev and Russel: Surgical margins in the veterinary cancer patient. Vet Comp Onc 15 (2017).

Kamstock et al: Recommended Guidelines for Submission, Trimming, Margin Evaluation, and Reporting of Tumor Biopsy Specimens in Veterinary Surgical Pathology. Vet Path 48 (2011).

Multiple sudden deaths were reported in a population of captive seahorses (Hippocampus abdominalis).

Histological examination revealed numerous ovoid to pear-shaped ciliated protozoal organisms measuring approximately 25×15 µm with a single nucleus. These were most numerous in subcutaneous fibrous connective tissue and muscle, but were also found within other tissues including the brain. The histological features were consistent with scuticociliates.

Protozoa are present in the periocular connective tissue and brain.

Background: Scuticociliates are free-living protozoa, causing disease in a wide range of marine fish species (1, 2). Often they act as opportunistic pathogens, but they have also been associated with mass mortality events, most recently in sea urchins (3). Environmental changes such as temperature fluctuations or poor water quality may predispose to infection.

Infected individuals may show lethargy, as well as areas of skin depigmentation or ulceration. Initially the organisms may be limited to the skin and gills but may then progress to systemic infections (4).

The infection may be diagnosed using wet mounts of skin or gills, but these may not yield sufficient protozoa, and often histopathology is required for diagnosis.

1. Di Cicco E,  Paradis E,  Stephen C,  Turba ME, and Rossi G. Scuticociliatid ciliate outbreak in Australian pot-bellied seahorse Hippocampus abdominalis (Lesson , 1827): Clinical signs, histopathologic findings and treatment with metronidazole Journal of Zoo and Wildlife Medicine44(2), 435-440, (1 June 2013). https://doi.org/10.1638/2012-127R1.1
2. Stidworthy MF, Garner MM, Bradway DS, et al. Systemic Scuticociliatosis (Philasterides dicentrarchi) in Sharks. Veterinary Pathology. 2014;51(3):628-632. doi:1177/0300985813492800
3. Hewson I et al. A scuticociliate causes mass mortality of Diadema antillarum in the Caribbean Sea. Sci Adv. 2023 Apr 21;9(16):eadg3200. doi: 10.1126/sciadv.adg3200
4. Edward J Noga. Fish Disease: Diagnosis and Treatment. 2^nd Wiley-Blackwell 2010. Chapter 8, pg. 141-143.