Introduction

Histoplasmosis is a common endemic mycosis caused by the thermally dimorphic fungus Histoplasma capsulatum which exists as a mold in the environment with conversion to a yeast at body temperature within the human host. Most infections result from inhalation of aerosolized microconidia from the environment, most commonly affecting the lung. Solid organ transplant (SOT) recipients and other immunocompromised hosts have a propensity for severe infection inclusive of extrapulmonary and disseminated disease given their inability to contain the infection, with significant associated morbidity and mortality. This review provides a general topic update, with emphasis on (1) the expanding geographic distribution of histoplasmosis; (2) the relationship between coronavirus disease 2019 (COVID-19) infection and histoplasmosis; (3) advances in fungal diagnostics; and (4) novel antifungal therapies, approved or in the pipeline.

Epidemiology and Risk Factors

Histoplasmosis is the most common endemic mycosis reported in the SOT population and is among the most common endemic mycoses in the United States (US) and throughout the world [1, 2•]. Three varieties of Histoplasma have been identified, though H. capsulatum var. capsulatum and H. capsulatum var. duboisii predominate among humans. The former variety is most common with the widest geographic distribution, whereas the latter is localized to Africa [3••, 4]. Importantly, genomic sequencing and phylogenetic analyses demonstrate marked diversity within the Histoplasma genus. The identification of multiple cryptic species with varying geography and virulence has led to modifications of Histoplasma taxonomy [5, 6].

The geographic distribution and overall burden of disease worldwide is grossly underestimated due to lack of access to available diagnostics and infection recognition, as well as insufficient surveillance and reporting [2•]. For example, histoplasmosis is not a nationally notifiable disease in the US. Prior to 2016, it was reportable in only 12 states and lacked a standardized case definition [7]. In addition, maps detailing the geographic distribution of histoplasmosis date as far back as the 1940s based on skin testing surveillance studies [2•]. Figure 1 reflects ongoing efforts to detail the expansion of histoplasmosis beyond previously defined high endemicity regions such as the Ohio and Mississippi River Valleys in the US, Central and South America, areas of Africa and Asia [3••]. The revised maps draw from many sources including autochthonous case reports and series, skin testing surveillance, and environmental sampling. Explanations proposed for the geographic expansion of histoplasmosis include improved diagnostics and clinical recognition, climate change, environmental disruption, and expanding at-risk hosts [2•, 3••].

Fig. 1
figure 1

Estimated areas of histoplasmosis worldwide. Legend: from reference [3••] under a Creative Commons Attribution 4.0 International License (http://creativecommons.org/licenses/by/4.0/)

Full size image

Histoplasmosis is uncommon among SOT recipients despite their increased susceptibility to infection. Prospective epidemiologic data from 23 transplant centers in the Transplant-Associated Infection Surveillance Network (TRANSNET) reported a 1-year cumulative incidence of histoplasmosis of 0.1% [1]. Transplant-associated histoplasmosis may occur de novo after environmental exposure, from reactivation of latent infection, or rarely via transmission from the donor. Although lung transplant recipients are at greater risk for acquiring other opportunistic fungi from the environment via the respiratory tract, such as Aspergillus spp., case series describe histoplasmosis most commonly after renal transplantation, likely as it reflects the largest transplant group worldwide [1, 8, 9].

Risks for post-transplant histoplasmosis include applied immunosuppression and environmental exposure. Human-to-human transmission has not been described. Histoplasma is commonly found in soil with high nitrogen content due to bird and bat drop**s from nearby trees, coops, or caves [10]. Outdoor recreational activities, particularly spelunking, are a known risk. A review of 105 outbreaks of histoplasmosis in the US spanning 1938 to 2013 demonstrated that nearly one-third were work-related involving occupations in construction, demolition, or maintenance. Exposure to birds, bats, or “drop**s” were reported in 86% [11]. Enhanced histoplasmosis surveillance in nine US states also identified handling of plants and trees and digging soil to be common potential exposures [12], though no obvious geographic or other exposure was identified in 22% of the cases. This point highlights the pitfalls of relying on a lack of relevant exposures to exclude histoplasmosis, as doing so could risk delayed or even missed diagnoses resulting in poor outcomes [13].

Clinical Manifestations

Clinical manifestations of histoplasmosis are highly variable and often non-specific making diagnosis challenging. Infection is often reported within the first 2 years following transplant, presumably due to higher levels of immunosuppression in this earlier transplant period. However, histoplasmosis can occur at any interval after transplant, having been reported as late as 20 to 30 years post-transplant [1, 8, 14••, 15, 16]. Infections occurring in the immediate post-transplant period, particularly the first month, should signal an investigation for donor-derived infection [8, 17••].

Signs and symptoms are dependent on the organ(s) involved and disease severity. Most infections present subacutely, with progressive disseminated infection representing the most common manifestation among SOT recipients, ranging from 64 to 81% in large multicenter evaluations [1, 8, 15]. Fevers, night sweats, malaise, cough, dyspnea, weight loss, hepatosplenomegaly, and other gastrointestinal symptoms are often present [10, 18, 19]. The most common sites of dissemination include bone marrow, lymph nodes, spleen, liver, gastrointestinal tract, skin, and central nervous system (CNS). Fungemia is common [8, 15, 20]. In the largest series of CNS histoplasmosis (n = 77), the most common findings included headaches (60%), altered mental status (42%), and focal neurologic deficits (30%) [21].

Common laboratory findings include pancytopenia, transaminitis, hyperbilirubinemia, elevated inflammatory markers (e.g., erythrocyte sedimentation rate, C-reactive protein, and ferritin), and hypercalcemia [10, 20, 22]. Histoplasmosis-associated hemophagocytic lymphohistiocytosis has also been rarely described in the SOT population [23]. Findings on chest imaging are varied, including isolated nodules, miliary infiltrates, multifocal consolidation, and cavitary lesions [10, 24]. Additional common radiologic abnormalities include intra-thoracic and abdominal lymphadenopathy, and hepatosplenomegaly [10, 18]. Computed tomography or magnetic resonance imaging in CNS disease often reveal mass lesions, meningeal enhancement, and/or infarcts though may be normal [21].

Diagnostic Strategies

Establishing the diagnosis of histoplasmosis in transplant recipients typically requires a multifaceted approach. The first step is suspecting the diagnosis by recognizing the various manifestations of progressive disseminated histoplasmosis, the most common presentation in SOT recipients. Table 1 summarizes available diagnostics for histoplasmosis. The diagnostic gold standard is growth of Histoplasma from clinical specimens. While definitive, cultures from blood and tissue may take up to 4 to 6 weeks to demonstrate growth, which may delay actionable clinical intervention [25]. Direct visualization of yeast forms with morphologic features consistent with Histoplasma from suspected sites of involvement is more timely and, unlike culture, can impact clinical decision making in real time. Though morphologic features may overlap with other fungi, careful attention to differences in yeast size, shape, budding, and capsular characteristics highly support the diagnosis of histoplasmosis in the right clinical context. Positive biopsy specimens are most commonly obtained from liver, lung, lymph nodes, bone marrow, and skin. Associated granuloma formation may also be seen, though less consistently in immunosuppressed hosts owing to the less robust host response to infection. With a high burden of infection, fungal forms within neutrophils may be identified on peripheral blood smear [26]. Cytopathologic examination of fine needle aspirate specimens from involved tissues may also be positive and provide a non-invasive, economical, and expedient diagnostic approach [27•]. The sensitivity of culture and direct visualization from histopathologic and cytopathic specimens is variable and dependent on the organism burden, site of involvement, and host factors [28•].

Table 1 Laboratory diagnosis for histoplasmosis [19, 21, 25, 26, 27•, 28•, 29,30,31,32,33,34, 36, 37, 44•, 45, 47, 48••]
Full size table

Prior to the development of more rapid diagnostic assays for histoplasmosis, acute and convalescent antibody detection was used to establish the diagnosis. The two available methods, testing by immunodiffusion and complement fixation, have differing sensitivity and specificity characteristics and are most useful in subacute and chronic forms of histoplasmosis [29]. However, in addition to the 4 to 8 weeks lag in antibody detection after acute infection, background seropositivity in individuals with prior exposure to endemic areas or past infection confound test interpretation. Serologic testing in immunosuppressed individuals is even more challenging as the effects of immunosuppressive therapy or underlying disease may blunt the antibody response to infection and decrease test sensitivity. Data demonstrate that only 25–30% of SOT recipients develop a serum antibody response [19]. However, CSF anti-Histoplasma antibody detection was found to be a useful adjunct to CSF antigen detection in diagnosing Histoplasma meningitis, even in immunosuppressed individuals [21, 30].

Owing to the challenges of culture identification and serologic testing, which are particularly problematic in transplant recipients and other immunocompromised hosts, antigen testing has become the primary modality for diagnosing histoplasmosis. The first antigen assay was introduced in 1986, with the development of newer iterations of enzyme immunoassays (EIAs) over time with quantitative capabilities and improved performance. Currently there are two commercially available EIAs. Most published data and clinical experience stems from use of the MiraVista® (MiraVista Diagnostics, Indianapolis, IN, USA) assay, which was the first assay developed, now in its third generation. The IMMY® (Immuno-Mycologics [IMMY], Norman, OK, USA) ALPHA EIA assay, now replaced with the subsequently developed IMMY® clarus Histoplasma galactomannan monoclonal EIA assay, has the advantage of the ability to be processed in local facilities rather than requiring shipment to a reference laboratory. These EIA assays demonstrate reasonable concordance, with overall sensitivity dependent on the type of specimen sent, the underlying burden of infection, and clinical manifestation [31,32,33,34].

Owing to the concentration of antigen in urine, urine testing is marginally more sensitive than serum antigen testing, with some data suggesting enhanced sensitivity with combined testing [35]. Antigen testing may also be performed on other body fluids, including bronchoalveolar lavage (BAL), CSF, and pleural fluid, though result interpretation is less established. The development of the quantitative component allows for serial monitoring to assess antigen clearance as a response to therapy, best supported in people living with human immunodeficiency virus (PLWH) [36].

Cross-reactivity with other fungal antigens is well demonstrated and limits EIA test interpretation, emphasizing the importance of interpreting the assay in the appropriate clinical context [32, 37]. False-positive BAL Histoplasma antigen results were found in 8% (5/60 patients) of pulmonary aspergillosis cases with high concentrations of Aspergillus galactomannan, though notably at low concentrations [38]. Conversely, the Aspergillus galactomannan assay was positive in 50% of serum and BAL samples from individuals with histoplasmosis, typically in those specimens with very high Histoplasma antigen concentrations. In this setting, the results could incorrectly lead to the conclusion there is co-infection and potentially impact treatment [39, 40]. The (1–3)-β-D-glucan assay as a screening tool for invasive fungal infection is widely used in clinical settings, but notably the assay does not have a US Food and Drug Administration (FDA)-approved indication for the diagnosis of histoplasmosis. Limited data demonstrate a sensitivity of serum (1–3)-β-D-glucan assay of 87 to 89% and specificity of 68% in disseminated histoplasmosis cases, with quantitative values correlating with urine antigen EIA concentrations [41, 42]. CSF (1–3)-β-D-glucan assay test performance was poor in Histoplasma meningitis cases [43].

While the development and broad availability of the Histoplasma antigen EIA assays have advanced the timely identification of histoplasmosis infections, logistical barriers related to the requirement of specialized laboratories, equipment, and personnel present challenges, particularly in resource limited areas. This recognition has led to the recent development of a qualitative lateral flow assay (LFA) for Histoplasma antigen detection. This assay has the advantage of using urine specimens at the point of care and providing results within 40 min of specimen collection without the need for specialized equipment. Direct cross comparison of results from urine specimens between the LFA and EIA assay has demonstrated high concordance between the two assays, with high test sensitivity in immunosuppressed individuals with a higher burden of infection [44•, 45]. However, both assays are limited by positive results due to cross-reactions with other fungal pathogens.

Currently, molecular detection of Histoplasma in the clinical setting is largely limited to fungal identification from culture isolates [46]. Matrix-assisted laser desorption ionization time-of-flight is increasingly applied in clinical microbiology laboratories for pathogen identification from culture isolates including both fungal morphologic stages of H. capsulatum [47]. While these approaches offer the advantage of rapid identification from primary cultures without needing a more prolonged and laborious method, they still require culture growth, which may require weeks to occur. Wide availability and applicability of molecular detection methods for H. capsulatum directly from clinical specimens remains elusive. Currently, there are no FDA-approved assays for detecting Histoplasma species directly from human specimens. Tissue-based polymerase chain reaction (PCR) testing and sequencing, along with broad-range PCR of fungal 28S ribosome, are available at reference laboratories, but are largely cost prohibitive for routine use. Available data are largely limited to case reports and small case series. Cross comparisons of studies are challenging due to the use of heterogeneous molecular assays using different molecular targets, multiple specimen sources, and various comparison assays [48••]. As with other diagnostic assays, the overall sensitivity of molecular techniques is superior with higher organism burden [49]. Performance evaluation and clinical validation of these assays require further refinement, but emerging data suggest molecular methods will play an increasing future role in our diagnostic armamentarium for histoplasmosis.

Management of Histoplasmosis

Most Histoplasma infections in immunocompetent individuals do not necessitate directed therapy [50••]. In contrast, SOT recipients can develop severe and disseminated infection warranting timely initiation of antifungal therapy and reduction of immunosuppression [14••]. While infections in SOT recipients are best categorized as mild, moderate, or severe, well-defined characteristics of severity in this population are lacking as are randomized controlled trials comparing the efficacy of antifungal therapies.

Available antifungal therapies used to treat histoplasmosis are detailed in Table 2; however, the mainstays of therapy have historically been amphotericin B (AmB) and/or itraconazole (ITZ). Intravenous AmB remains the initial drug of choice for severe infection. Liposomal AmB is preferred over conventional AmB deoxycholate (AmB-d) due to its more favorable safety and tolerability profile and improved efficacy as demonstrated in other immunocompromised populations such as PLWH [51]. The American Society of Transplantation (AST) guidelines recommend step down to oral ITZ therapy following at least 1 to 2 weeks of AmB alongside evidence of clinical stabilization [14••]. For patients with CNS disease, longer courses of AmB therapy (i.e., 4 to 6 weeks) are recommended [50••, 52•]. Mild to moderate disease can be treated with ITZ monotherapy. In general, antifungal therapy should be continued for a minimum of 12 months irrespective of initial disease severity. Indefinite therapy may be considered in patients necessitating continued high-level immunosuppression or after relapsed disease, particularly with CNS involvement [53].

Table 2 Antifungal therapy for histoplasmosis [50••, 54, 55, 57, 67••, 92,93,94,95,96,97,98,99,100]
Full size table

Conventional ITZ (C-ITZ) is available as both an oral capsule and solution [54, 55]. C-ITZ capsules have variable bioavailability, which is particularly poor when administered with concomitant acid-suppressive therapies. Absorption can be optimized when C-ITZ is taken with a fatty meal and/or acidic drink. Conversely C-ITZ solution is minimally impacted by acid-suppressive therapies but requires administration on an empty stomach. C-ITZ solution has generally been preferred owing to its superior bioavailability though palatability can be problematic. Therapeutic drug monitoring is essential with both formulations to ensure adequate exposure [56]. Super bioavailable itraconazole (SUBA-ITZ) is a novel formulation of ITZ developed to address issues of tolerability and bioavailability observed with C-ITZ formulations [57]. It is available as an oral capsule formulation and applies a solid dispersion of ITZ in a polymetric matrix. Studies of SUBA-ITZ in healthy adults demonstrated no significant reductions in bioavailability in the presence of acid-suppressive therapies or in fed versus fasted states [58]. A compilation of seven crossover studies in healthy adults receiving SUBA-ITZ and C-ITZ capsules demonstrated improved bioavailability of SUBA-ITZ (173%) compared to C-ITZ capsules with less variability in ITZ exposure [59]. In addition, steady-state findings from an open-label crossover pharmacokinetic analysis of SUBA-ITZ and C-ITZ capsules showed greater attainment of therapeutic ITZ trough values (i.e., > 1000 ng/mL) in the SUBA-ITZ than C-ITZ group (81 versus 44%, respectively) [60]. SUBA-ITZ was approved by the FDA in 2018 for the treatment of specific fungal infections including histoplasmosis. Published results from a Mycosis Study Group-sponsored phase 3 multicenter, randomized, open-label study of invasive endemic fungal infections evaluating the pharmacokinetics, safety, efficacy, tolerability, and health economics of oral SUBA-ITZ compared to C-ITZ capsules are highly anticipated (NCT03572049).

Echinocandins are not currently recommended for histoplasmosis treatment due to a lack of supportive clinical data. However, fluconazole (FCZ) has been used as an alternative to ITZ in settings such as intolerance. Concerns with application of FCZ for Histoplasma include reduced in vitro activity, delayed fungemia clearance, and resistance emergence [50••, 61, 62]. Newer-generation extended-spectrum azoles including voriconazole (VCZ), posaconazole (PCZ), and isavuconazole (ISZ) may be suitable alternatives to ITZ though data are limited to case reports and series [63,64,65,66]. A particular advantage may exist in CNS infection, as VCZ has improved CNS penetration compared to ITZ but necessitates further study. Hendrix et al. published the only data to date comparing outcomes with VCZ and ITZ for histoplasmosis in a single-center, retrospective evaluation in 194 cases that included 21 (10.8%) transplant recipients [67••]. Survival analysis in patients receiving ITZ (90.2%) and VCZ (9.8%), either as primary or step-down therapy following AmB, demonstrated a statistically significant association of VCZ therapy with early mortality during the first 6 weeks (hazard ratio, 4.3; 95% confidence interval, 1.3–13.9; P = 0.015). Results of therapeutic drug monitoring were not reported and only one case with CNS disease was treated with VCZ. Potential explanations for poorer survival with VCZ include suboptimal drug exposure and resistance emergence. As previously mentioned, resistance has been demonstrated with FCZ and is due to a point mutation in cytochrome P450 enzyme 14α-demethylase (CYP51p) [68]. This mutation results in increased minimal inhibitor concentrations (MIC) for both FCZ and VCZ but does not appear to impact the MIC for the other extended-spectrum azoles [68, 69]. Despite the limitations of this retrospective evaluation, caution is advised when utilizing VCZ for histoplasmosis, particularly in the setting of preceding FCZ exposure. In situations of ITZ intolerance, the authors prefer PCZ based on resistance concerns and improved tolerability despite the limited clinical data. Prospective controlled trials are necessary to determine whether VCZ and other extended-spectrum azoles are effective therapeutic options for histoplasmosis, including CNS disease, and to further understand the potential for resistance emergence.

There are additional important considerations when managing histoplasmosis in the SOT recipient. Drug toxicities and drug interactions, especially those related to the cytochrome P450 isoenzyme system, remain a significant issue with azole therapies (see Table 2). Polypharmacy is common in the transplanted host and often includes other narrow therapeutic index agents such as calcineurin inhibitors (CNIs) impacting the CYP450 isoenzyme system. Thus, azole use necessitates careful dose adjustment of relevant medications and close monitoring. Reducing immunosuppression during acute infection is also paramount. However, an optimal strategy is lacking, with decisions typically individualized to the transplant recipient based on the transplant type and timeline, status of allograft rejection, and net state of immunosuppression. Mycophenolate has been identified as a risk factor for severe disease and failure to reduce CNI exposure has been associated with relapsed disease; hence, reductions in these agents should be prioritized [8]. While immune reconstitution syndrome has been described during treatment of disseminated histoplasmosis, it is uncommon in SOT and other immunocompromised populations such as PLWH [70, 71]. Finally, monitoring of urine and serum Histoplasma antigen levels during and after therapy is recommended to assess response to infection and monitor for relapse. The Infectious Diseases Society of America (IDSA) and AST guidelines recommend monitoring antigen levels at regular intervals, including at therapy initiation, 2 weeks, 1 month, and approximately every 3 months thereafter during therapy and for at least 6 to 12 months post-cessation [14••, 50••]. Additional testing should be performed as clinically indicated, such as scenarios when there is concern for relapse or with marked augmentation of immunosuppressive therapy (e.g., re-transplantation and severe allograft rejection). Not all SOT recipients will completely clear antigenemia and/or antigenuria at the end of appropriate therapy; however, most can still safely stop antifungal therapy without relapse, assuming all other indicators support clinical response [15, 72]. Of note, quantitative antigen values may vary depending on the specific EIA assay applied; hence, a consistent laboratory should be utilized for serial monitoring when feasible.

Novel Fungal Therapies for Histoplasmosis

There are additional drugs either in the antifungal pipeline or FDA-approved for other indications that have Histoplasma activity. These agents may ultimately supplement the limited fungal armamentarium for histoplasmosis and other endemic mycoses. For example, new polyene formulations in development include encochleated amphotericin B deoxycholate (MAT2203; Matinas BioPharma Nanotechnologies, Bedminster, NJ) which utilizes a novel lipid nanocrystal to allow for oral administration. This therapy lacks many of the adverse effects associated with intravenous AmB including infusion reactions, nephrotoxicity, and electrolyte disarray [73, 74]. Tetrazoles are next-generation azoles in development to abrogate management issues such as drug interactions given their greater selectivity for fungal Cyp51 compared with the mammalian cytochrome P450 enzyme system [75, 76]. Among these, VT-1598 (Viamet Pharmaceuticals, Durham, NC) has a broad spectrum of activity including Histoplasma and was granted orphan drug designation for the treatment of coccidioidomycosis. Ibrexafungerp is an oral triterpenoid antifungal with a mechanism of action similar to echinocandins, inhibiting 1,3-β-D-glucan synthase [75, 77, 78]. It received FDA approval in 2021 for treatment of vulvovaginal candidiasis. An ongoing phase 3 multicenter, open-label study evaluating the efficacy and safety of ibrexafungerp in patients with fungal diseases refractory to or intolerant of standard treatment was expanded to include histoplasmosis (FURI, NCT03059992) [77]. Other novel agents with Histoplasma activity include olorofim, fosmanogepix, and nikkomycin Z [76, 78, 79•, 80,81,82,83]. Both olorofim and fosmanogepix have activity against Histoplasma, are available in oral and intravenous formulations, and are actively being pursued for multiple fungal applications. Olorofim, an antifungal in the orotomide class, inhibits dihydroorotate dehydrogenase in the pyrimidine biosynthesis pathway [81]. Fosmanogepix, the oral prodrug of manogepix, inhibits the fungal enzyme Gwt1 important for mannoprotein adherence to the cell wall during establishment of infection [79•]. Finally, nikkomycin Z is a chitin synthase inhibitor and impacts chitin production, an important structural component of fungal cell walls. It demonstrates activity against histoplasmosis both as monotherapy and in combination with FCZ [83]. Renewed interest in nikkomycin Z for coccidioidomycosis treatment alongside the development of an extended-release formulation to offset its short half-life may hold promise in future application for endemic mycoses including histoplasmosis [84].

COVID-19 and Histoplasmosis

Emerging reports have described a temporal relationship between COVID-19 infection and invasive fungal infections, including histoplasmosis [85•]. Reports of histoplasmosis develo** during and after COVID-19 infection are limited to small series and case reports, occurring among both immunocompromised and immunocompetent individuals [86,87,88,89,90]. Only one such case has been reported in a kidney transplant recipient, though additional anecdotal cases in SOT recipients have occurred (author personal communication) [90].

There are two main proposed mechanisms to explain an association between histoplasmosis and COVID-19 infection. First, parenchymal lung injury related to COVID-19 infection may predispose individuals to develo** acute pulmonary histoplasmosis after an inhalation of environmental Histoplasma conidia. Second, SARS-CoV-2-associated immune modulation or that related to corticosteroid therapy used for COVID-19 infection management may lead to reactivation of latent Histoplasma infection in those individuals previously infected. Further, these factors may act synergistically in an infected individual. In many of the reported cases, interpretation of the diagnostic testing in the clinical context is challenging such that firm conclusions regarding a true link between Histoplasma and COVID-19 infections are lacking. More data are needed, but these early cases suggest clinicians should maintain heightened clinical suspicion for histoplasmosis, especially in endemic areas, among patients with concurrent or recent COVID-19 infection.

Peri-transplant Donor and Recipient Considerations

Recipient Evaluation

General consensus based on data compiled over 2 decades support that pre-transplant screening for histoplasmosis in transplant candidates is not indicated, even for those in endemic areas. Early support for this approach came from a retrospective review of 449 SOT recipients from a hyperendemic region, where approximately one quarter of SOT candidates had either a positive Histoplasma serology or chest radiograph evidence of prior infection [91]. None developed post-transplant histoplasmosis despite no antifungal prophylaxis, suggesting the risk for reactivation infection is negligible in this setting. Transplant candidates with active histoplasmosis should be deferred at least until there is significant clinical improvement, or if feasible, until therapy is completed. Active histoplasmosis within the prior 2 years may be a basis for antifungal prophylaxis after transplantation, though the duration is not established [50••]. Subsequent case series have further supported this approach, with targeted post-transplant antifungal prophylaxis for other selected recipients [8, 15, 19, 72].

Donor Evaluation

Given the low incidence of histoplasmosis in healthy donors and rarity of reported donor transmission events, even in endemic areas, routine donor screening is not indicated. However, if the donor demonstrates clinical findings such as non-hemorrhagic neurological disease, unexplained fever, and/or pneumonia, histoplasmosis should be considered and testing performed. Though procurement may proceed before tests result, organs should be carefully inspected for findings suspicious for histoplasmosis, such as hepatosplenomegaly, granuloma formation, and lymphadenopathy. In addition to obtaining tissue specimens for histopathology and culture, serology and antigen EIA testing should be performed. The collective results will inform recipient management following allograft implantation. Potential living donors with active histoplasmosis should be deferred and treated with antifungal therapy at least 3–6 months prior to consideration for organ donation [17••].

Management Recommendations

The optimal management approach for clinical scenarios where is there is concern for possible donor Histoplasma transmission and pre-existing recipient infection remain undefined. Accordingly, there exists significant practice variability among individual transplant centers, geographic regions, and organ transplanted. Recognizing the paucity of data, in 2012, the AST published guidance [17••]. Per these recommendations, azole prophylaxis should be offered if the donor tissue fungal stains are positive but cultures and antigen assays are negative and/or the donor complement fixation serology is ≥ 1:32. The recommended prophylaxis duration is 3 to 6 months. For lower risk scenarios, observation without azole prophylaxis with antigen EIA monitoring every 3 months is a reasonable approach. If active histoplasmosis is confirmed in the donor after allograft implantation, the recipient should be treated with azole therapy for at least 1-year post-transplant.

While appreciating the attempt to provide a construct for uniform practice across the transplant community, these recommendations were based largely on expert opinion and now date back a decade, leaving leeway for alternative present day approaches. Without updated data, testing and management of donors and recipients remain variable and individualized based on the infection burden and underlying recipient risk. Lung transplant recipients are considered higher risk for develo** active histoplasmosis based on requirements for relatively higher baseline immunosuppression compared to other transplant groups as well as the pulmonary reservoir of latent or active Histoplasma organisms. Newer, extended-spectrum azoles with improved tolerability also introduce expanded options for prophylaxis and treatment. It is important to recognize that histoplasmosis remains a rare infection (< 1%) in the post-transplant setting, even in endemic areas [1].

Conclusions

Though an uncommon infection in the SOT population, histoplasmosis has significant clinical implications. Given the myriad of clinical manifestations associated with infection in SOT recipients and the expanding worldwide distribution, histoplasmosis should increasingly be suspected in the appropriate clinical context. Laboratory testing continues to evolve to optimize timely diagnosis and monitoring and the antifungal armamentarium is growing. With diligent monitoring for active infection and targeted post-transplant antifungal prophylaxis, serious infection can largely be averted in the highest risk transplant recipients.