OBM Transplantation

(ISSN 2577-5820)

OBM Transplantation is an international peer-reviewed Open Access journal, which covers all evidence-based scientific studies related to transplantation, including: transplantation procedures and the maintenance of transplanted tissues or organs; assimilation of grafted tissue and the reconstitution of removed organs or parts of organs; transplantation of heart, lung, kidney, liver, pancreatic islets and bone marrow, etc. Areas related to clinical and experimental transplantation are also of interest.

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Open Access Review
Non-CMV Viral Infections Following Solid-Organ Transplantation – Focus on Human T-Cell Lymphotropic Virus Type-1 and Human Herpesviruses-6,-7 and -8

Katerina G. Oikonomou , Sarah Taimur *

Department of Medicine, Division of Infectious Diseases, Icahn School of Medicine at Mount Sinai, New York, NY, USA

Correspondence: Sarah Taimur

Academic Editor: Maricar Malinis

Special Issue: Diagnosis and Management of Infections in Solid Organ Transplant Recipients

Received: February 14, 2019 | Accepted: June 06, 2019 | Published: June 17, 2019

OBM Transplantation 2019, Volume 3, Issue 2, doi:10.21926/obm.transplant.1902066

Recommended citation: Oikonomou KG, Taimur S. Non-CMV Viral Infections Following Solid-Organ Transplantation – Focus on Human T-Cell Lymphotropic Virus Type-1 and Human Herpesviruses-6,-7 and -8. OBM Transplantation 2019;3(2):26; doi:10.21926/obm.transplant.1902066.

© 2019 by the authors. This is an open access article distributed under the conditions of the Creative Commons by Attribution License, which permits unrestricted use, distribution, and reproduction in any medium or format, provided the original work is correctly cited.

Abstract

In non-endemic regions of the world, human T-cell lymphotropic virus type-1 (HTLV-1) is an uncommon pathogen in the transplant host, but can be associated with significant morbidity and mortality. Careful assessment for risk factors, targeted screening and heightened awareness of the clinical presentation of HTLV-1 associated disease is necessary for timely recognition and management in the transplant host. The use of antiretroviral agents in the management of symptomatic disease due to HTLV-1 remains controversial. Human herpesvirus-6 (HHV-6) has long been recognized as a pathogen in the transplant host however, establishing pathogenicity remains a challenge in clinical situations. Chromosomally integrated HHV-6 has been reported in ~1% of the solid-organ and allogeneic stem cell transplant population; and is often mistaken for active infection. Increased recognition of this entity is needed to avoid unnecessary use of antiviral medications. Current guidelines recommend against screening and treatment of asymptomatic HHV-6 infection in the solid-organ transplant host. Human herpesvirus-7 (HHV-7) is often diagnosed as co-infection with other beta-herpesviruses, but pathogenicity is less clear. There continues to be no clinical syndrome solely attributable to HHV-7. Human herpesvirus-8 (HHV-8) infection following organ transplantation can be due to primary acquisition from donor or non-donor derived exposures; or secondary to reactivation of latent infection in a seropositive recipient. Kaposi sarcoma is the most common HHV-8 associated post-transplant complication however, there is increasing recognition of non- neoplastic syndromes of febrile illness with bone marrow suppression and hemophagocytic syndrome. Lack of standardized laboratory assays for HHV-8 remains an impediment to targeted screening of high risk organ donors and recipients. A multi-disciplinary approach is needed for management of HHV-8 associated diseases.

Keywords

HTLV-1; HHV-6; HHV-7; HHV-8; solid-organ transplantation

1. Introduction

Human herpesvirus-6,-8 (HHV-6,-8) and human T-lymphotropic virus type-1 (HTLV-1) are associated with significant morbidity and mortality in the solid-organ transplant host. The role of HHV-7 as a sole pathogen remains unclear; but co-infection with other beta-herpesviruses is frequently encountered in the immunosuppressed host. There are unique challenges associated with each of these viruses as it pertains to the organ transplant recipient. In case of HHV-6 and HHV-7, it is their ubiquitous nature, and frequent clustering of infection with Cytomegalovirus, that continue to pose diagnostic and therapeutic challenges to the transplant practioner. HHV-8 and HTLV-1 on the other hand, are geographically uneven viruses; and while well-established pathogens in the transplant host, need greater recognition in non-endemic regions of the world. This review aims to enhance our understanding of the impact of HHV-6,-7,-8 and HTLV-1 on the solid-organ transplant host; and highlights the diagnostic and therapeutic challenges that continue to be associated with these infections.

2. Human T-Lymphotropic Virus Type-1

Human T-lymphotropic virus type-1 (HTLV-1) is an RNA retrovirus that was first described in the 1980s [1,2]. An estimated 15 to 20 million individuals worldwide are infected with HTLV-1; which is endemic in parts of South America, the Caribbean, West Africa and Asia [3,4,5,6]. Based on data in blood donors, seroprevelance in the United States is less than 1% [7]. The most common mode of transmission of HTLV-1 in endemic areas is mother to child, most commonly through breast feeding [8]. Other modes of transmission include sexual intercourse, transfusion of cellular blood products and organ transplantation [9].

Following initial infection, HTLV-1 establishes a state of life-long latency in the lymphocytes. Most people remain asymptomatic but in an estimated 2%-5% of cases, symptomatic disease as HTLV-1 associated myelopathy tropical spastic paraparesis (HAM/TSP) or adult T-cell leukemia/lymphoma (ATL) can develop [4,10]. HAM/TSP usually presents with gait disturbances, lower extremity weakness, urinary incontinence, decreased sensation and upper motor neuron signs [11]. Some of the clinical manifestations of ATL are generalized lymphadenopathy, hypercalcemia and bone lesions.

2.1 HTLV-1 in Solid-Organ Transplantation: An Uncommon But Potent Pathogen    

While transmission through organ transplantation has been reported, there are only a few known cases of proven donor-derived HTLV-1 infection [12,13]. The rate of HTLV-1 associated disease appears to be even lower in the transplant host [11,13,14,15,16]; and this is evident in Organ Procurement Transplantation Network (OPTN)/United Network of Organ Sharing (UNOS) data that identified no cases of HTLV-1 infection among 162 recipients of 134 reactive HTLV-1/2-positive donors between 1999 and 2008 [17,18,19].

In SOT recipients, the clinical course of HTLV-1 disease can be different with shorter latency period and more rapid progression of disease [20,21]. The natural history of HTLV-1 in the transplant host however, is not well understood due to lack of long-term cohort studies. The factors that promote disease in SOT patients are an increased pro-viral load, immunosuppression and association with host human leukocyte antigen (HLA) subtypes (B-5401, DRB1-0101). HTLV-1 antibodies usually develop 1-3 months after exposure and are positive lifelong [10,12,22,23].

2.2 HTLV-1 - Pathogenesis and Diagnosis

The first step in the diagnosis of HTLV-1 infection is screening with enzyme immunoassays (EIA). Positive screening tests can then be confirmed using line immunoassays or Western blot [24]. Currently available screening assays have high sensitivity but do not differentiate between HTLV-1 and 2; and have low positive predictive value in areas of low seroprevelance such as the United States. HTLV-1 is a highly cell bound with only small amounts of free virus present in the circulation. The virus infects lymphocytes and the pro-virus undergoes integration into the host genome. PCR testing for viral RNA on non-cellular specimens such as plasma is therefore of limited diagnostic utility [25]. Using PCR for measurement of HTLV-1 pro-viral DNA load in peripheral blood mononuclear cells appears to be a promising tool that may have a role in diagnosis of cases (particularly when standard confirmatory assays are indeterminate) and to assess risk of HTLV-1 associated disease [26,27,28]. This is an encouraging development but further research is needed to better understand the diagnostic role of pro-viral DNA measurement in the clinical care of patients.

2.3 Finding the Balance - Shift away from Universal Donor Screening

In light of low positive predictive value of available testing and concern about preclusion of healthy donor organs based on false positive results, the policy of universal donor screening for HTLV-1/2 was put on the table for discussion. These concerns were further supported by analysis of universal HTLV-1/2 screening of deceased donors which revealed that the ratio of false positive to true positive screening was 40:1 [4,29]; and that an estimated 167–227 uninfected organs were rejected each year due to false positive screening results [4,17].

In 2009 mandatory, universal donor screening for HTLV-1, was eliminated by the OPTN/UNOS. However, further recommendations were made by the Organ Procurement Transplantation Network Ad Hoc Disease Transmission Advisory Committee (DTAC) that targeted screening of donors presumed to be at high risk for HTLV-1 infection should continue, and all positive donor screening tests should be confirmed retrospectively to allow appropriate follow up of the recipients [17]. The American Society of Transplantation Infectious Disease Community of Practice (AST IDCOP) also recommends targeted screening of high risk donors and recipients [4]. Since organs from donors who screen positive for HTLV-1 can be used for transplantation, it is essential that the recipient is informed about the potential risks as part of the informed consent process required for transplantation. The optimal management and follow up of recipients receiving organs from donors proven or suspected to have HTLV-1 is unknown, and there are currently no established guidelines. The AST IDCOP recommends using a combination of laboratory tests and clinical monitoring for follow up of recipients with confirmed or high suspicion of donor infection. Transplant practioners must also be educated on clinical manifestations of HAM/TSP and ATL, and must maintain a heightened awareness in follow up of high risk organ recipients for timely diagnosis and management of HTLV-1 associated complications.

2.4 HTLV-1 Associated Disease: HAM/TSP and ATL

The diagnosis of HAM/TSP can be particularly challenging and transplant practioners should be aware of the WHO criteria for timely recognition and appropriate work up [30]. Diagnosis can be missed or delayed due to rarity of the HTLV-1 infection outside of endemic areas, lack of pre-transplant serologic screening and low frequency of associated disease in infected SOT recipients. In patients developing ATL the treatment consists of chemotherapy; and steroids, interferon-alpha, and immunoglobulin therapy might be employed in cases of HAM/TSP. While sometimes employed, the role of antiviral medications in HTLV-1 associated disease remains controversial.

Antiretroviral medications including zidovudine and raltegravir have been shown to have in-vitro activity against HTLV-1 [31]; however the in-vivo results have been disappointing [32,33,34]. This lack of concordance is explained by the lack of continuous viral replication cycles in chronic carriers of the infection [4,34,35]. Limited clinical data suggest efficacy of prosultiamine with improved motor strength, urinary bladder function and pro-viral DNA in patients with HAM/TSP [35,36]. Prosultiamine is a thiamine derivative has been found to have activity against HTLV-1 infected cells by inducing apoptosis [37]. While promising, these findings need investigation in larger randomized clinical trials to better understand the benefit to patients. Based on expert opinion, it has been suggested that post-exposure prophylaxis may be beneficial to prevent HTLV-1 infection following exposure through organ transplantation. There is however no data on the efficacy of this approach and it is currently not approved or recommended by the CDC [3].

3. Human Herpesvirus-6

Human herpesvirus-6 (HHV-6) is a lymphotropic beta (β)-herpesvirus, which was first isolated from peripheral blood lymphocytes of immunosuppressed patients in 1986 [38]. Initially classified as variants of the same virus, HHV-6A and B are now recognized as separate species [39]. In the vast majority of the population, HHV-6 exposure occurs during early childhood, and initial infection is followed by establishment of life long latency inside the mononuclear cells [40,41,42].  Majority of human infections are due to HHV-6B with an estimated prevalence of 97% [43]. In addition to blood mononuclear cells, a wide variety of body tissues are infected with HHV-6 including salivary glands, tonsils, brain, liver and kidney [44]. In a smaller percentage of the population, HHV-6 viral genome undergoes integration into the host chromosome [45,46,47]. The ability to establish latency through integration into the host genome is a unique property of the HHV-6 virus. This entity is termed as chromosomally integrated HHV-6 (CIVVH-6), and is estimated to occur is 1% of the human population worldwide [46]. The clinical significance and diagnosis of CIHHV-6 in the transplant host will be discussed in detail later in this manuscript.

Since the vast majority of adults are seropositive, HHV-6 infection in the post-transplant setting most commonly is due to reactivation of endogenous virus [48,49]. Primary infection through transmission from the community or the allograft itself has been reported but very uncommon [50,51]. HHV-6B has been implicated as the cause of majority of infections in adult SOT recipients [39,43,52,53]; with a much smaller percentage due to HHV-6A.

3.1 Clinical Manifestations of HHV-6 in the Solid-Organ Transplant Host: Direct and Indirect Effects

The reported incidence of symptomatic infection in the SOT recipients varies between clinical series but overall despite high seroprevelance, the incidence of symptomatic HHV-6 infection following organ transplantation is low [48,49,54]. Most HHV-6 infections represent reactivation of latent virus in the early post-transplant period (first 4-6 weeks) with fewer reports of late onset infections (months to years) [48,55].

Clinical manifestations of HHV-6 can be attributed to direct and indirect effects. Among the direct effects, the most commonly reported include febrile illness with bone marrow suppression and encephalitis [56,57]. Other reported manifestations are gastrointestinal (gastroduodenitis, colitis), pneumonitis, fulminant hepatic failure. Reported manifestations of HHV-6A in the transplant population include giant-cell hepatitis [58], hemophagocytic syndrome [50], and hepatitis with pancytopenia [51]. There are indirect effects of HHV-6 due to immunomodulatory properties which include recurrence of hepatitis C infection [59,60], Cytomegalovirus (CMV) infection [61,62,63], allograft rejection [62,64], and fungal infections [54,62,65]. The increased risk of CMV infection/disease in association with HHV-6 was clearly shown in a study by Kumar et al where risk of CMV disease increased by 3.59-fold in patients with HHV-6 infection (95% CI, 1.53-8.44; P=0.003) [54]. Limited data is suggestive of an association between HHV-6 and bronchiolitis obliterans syndrome but further research is warranted to better understand and validate these findings [66].

There are many reports of HHV-6 and CMV co-infection in SOT recipients. On genomic analysis, HHV-6 is most closely related to human CMV with 66% similarity in amino acid sequences [67]. In a study of liver transplant recipients who were found to be positive for both viruses, HHV-6 was on average activated earlier than CMV in most patients (HHV-6 activation: 19.8 ± 4.5 days vs. CMV activation: 28.4 ± 60.5 days - P = 0.001), with a 9-day time gap between the activation of the two viruses. Regression analysis of the time of CMV and HHV-6 activation revealed a linear relationship [68].

3.2 Diagnosis of HHV-6: Establishing Pathogenicity of a Ubiquitous Organism

Over the years, HHV-6 has been recognized as a pathogen in the transplant host. However, establishing pathogenicity of HHV-6 in clinical situations remains a challenge. Clinical manifestations of HHV-6 overlap with other herpes viruses, particularly CMV; and it is not uncommon to find both viruses in the work up of patients with suspected infection. In these scenarios, HHV-6 is often questioned as the true-pathogen in comparison to CMV. Other features of HHV-6 which make the diagnosis of active infection challenging, are the ubiquitous nature of the virus, high seroprevelance in humans, establishment of clinical latency and integration of viral genome into the human chromosome. Careful consideration must be given to the intrinsic properties of HHV-6 including CIHVV-6, and the limitations of current diagnostics to avoid unnecessary antiviral therapy [53].

Diagnostic approach to HHV-6 has been characterized broadly into indirect and direct [44,69]. Indirect diagnostics are based on detection of HHV-6 antibodies, while direct diagnostics employ methods for detection of whole virions or viral components and include viral culture, HHV-6 antigen detection, and PCR.

3.2.1 Serologic Testing

Serology for HHV-6 is useful as a screening tool to determine risk of future infection in an immunosuppressed host but since seroprevelance in the adult population is high, it’s utility in the diagnosis of active infection is limited. Furthermore, currently available serologic tests are species non-specific hence unable to differentiate between HHV-6A and B species and may have cross-reactivity with other β-herpes viruses [44,69].

3.2.2 Polymerase Chain Reaction

Most commonly employed direct diagnostic testing is based on detection of viral nucleic acids by PCR. PCR testing for HHV-6 is commercially available for a broad range of clinical specimens and does provide differentiation between HHV-6A and B species however, standardization of testing methodology and measurement units is lacking [44,69]. Active infection is often diagnosed based on quantitative PCR testing (Q-PCR) but there are no standardized thresholds for interpretation of active infection versus latent versus CIHHV-6. For diagnosis of active infection, reverse-transcriptase PCR can be employed that can differentiate between latent and actively replicating virus by detection of viral transcripts. This technique was described in 1999 where viral transcripts were detected by amplification of mRNA corresponding to major structural antigens in peripheral blood mononuclear cell samples taken from children with recent clinical illness suggestive of HHV-6. In comparison with DNA-PCR and viral culture performed on the same specimens, reverse-transcriptase PCR was found to have sensitivity of 95% and specificity of 98.8% [70].

3.2.3 Viral Antigen Based Testing

With the advent of Q-PCR testing for HHV-6, the role of antigenemia assays as screening tools for detected of viremia is limited. However, detection of viral antigens in tissue using monoclonal antibodies and immunohistochemical testing is still of benefit for the diagnosis of tissue invasive disease [55,71].

3.2.4 Plasma versus Whole Blood PCR

Q-PCR testing for diagnosis of HHV-6 infection is most often performed on non-cellular specimens such as plasma despite the fact that the virus is highly cell associated. This has raised questions about the adequacy of plasma as a sample for HHV-6 testing. In fact, recent data revealed that the source of HHV-6 DNA detected in plasma infact corresponds with lysis of blood cells rather than active virus production due to infection of lymphoid tissue or organs [72]. Further research on patients samples with various stages of HHV-6 infection (primary, past, congenital, no infection, CIHHV-6) revealed that in comparison with virus isolation by culture, plasma HHV-6 real time-PCR had a sensitivity of 92% for identification of primary infection and specificity of 84% [73]. All patients in this study with CIHHV-6 had detectable virus by plasma HHV-6 real time PCR but only 5.6% had detection of active viral transcripts by reverse-transcriptase PCR [73]. This data suggests that Q-PCR testing on plasma is not a reliable marker of active infection and in order to distinguish active infection from CIHHV-6, testing on whole blood might be more specific [73,74]. Q- PCR testing on whole blood is available commercially but data are limited. As with testing on plasma, there are no standardized cut-offs for interpretation of active versus latent infection.

It is important to recognize the differences between the currently available diagnostic tests for HHV-6, to choose the test appropriate for the clinical question at hand. Clinicians must also bear in mind, that current testing for HHV-6 is limited not only by the lack of clinically meaningful cut-offs for active infection, but also by the lack standardized WHO units of IU/mL; contributing to inter-laboratory variation in results. We make the following recommendations in this regard.

  • Serologic testing is not of utility in the diagnosis of active infections. A positive serology with negative Q-PCR testing indicates latent HHV-6 infection.
  • In a patient with clinical signs and symptoms attributable to HHV-6, positive Q-PCR testing can be indicative of active infection however; important to remember that Q-PCR testing does not distinguish CIHHV-6 from active viral replication.
  • CIHHV-6 is typically an asymptomatic patient with a very high HHV-6 viral load. To distinguish CIHHV-6 from active infection, reverse-transcriptase PCR testing can be employed.
  • The current role of viral antigen based tests in the diagnosis of active HHV-6 infection is largely limited to immunohistochemistry for detection of viral antigen in tissue specimens.

3.3 Chromosomally Integrated HHV-6: Clinical Significance and Diagnosis

Based on the work of multiple investigators, CIHHV-6 is defined as the presence of ≥1 HHV-6 DNA copies/cell [46,75,76]. Clinical significance of CIHHV-6 in the immunosuppressed host is an area of active research. The prevalence of CIHHV-6 in the solid-organ and allogeneic stem cell transplantation appears to be low (~ 1%) [77]. Study of the integrated HHV-6 viral genome has shown that these viruses differ greatly from their non-integrated counterparts. These integrated genomes however, in majority (CIHHV-6B 95% and CIHHV-A 72%), do contain a complete complement of intact viral genes and appear to be capable of reactivation from the latent state [78]. While the clinical significance of CIHHV-6 in the organ transplant host remains undefined, limited data suggest that the rate of clinical disease due to CIHHV-6 is low [53]. A review by Lee et al of 9 SOT patients with CIHHV-6 [53], showed that the median HHV-6 DNA copies per ml of whole blood in LT recipients was 2.7 x 106 (1.1 x 106 - 18.3 x 106) and in kidney transplant patients was 5.1 x 106 genomes per ml of whole blood. In the same series, a small bowel recipient had HHV-6 DNA PCR done on hair follicles which revealed DNA concentration of 11 x 106/106 cells, consistent with CIHHV-6 [76]. None of these patients developed clinical disease due to HHV-6 infection and 2 patients were treated with antiviral agents for mistaken diagnosis of active infection [53,79]. heightened awareness of this entity is therefore needed and a suggested approach by experts is that in patients with suspected CIHHV-6 (typically DNA load of > 5.5 log10 copies per ml of whole blood), reverse-transcriptase PCR be considered to rule out active infection due to HHV-6 and avoid unnecessary use of antiviral agents [47,73].

3.4 Treatment of HHV-6: Indications for Antiviral Therapy

Acyclovir and valacyclovir have no significant activity against HHV-6 at doses used in patient care. It is however susceptible to ganciclovir, foscarnet and cidofovir [44,80], at doses similar to those used for treatment of CMV. HHV-6 genes which are of relevance to antiviral treatment include U69 and U38. U69 encodes a protein kinase, which is involved in the initial phosphorylation of ganciclovir that is subsequently tri-phosphorylated to its active form by cellular kinases. U38 encodes the viral DNA polymerase [80] which is the site of action of ganciclovir, foscarnet and cidofovir.

Successful use of ganciclovir and foscarnet for symptomatic HHV-6 infection has been reported in transplant recipients, but no controlled trials have been performed to prove efficacy [81,82,83,84]. A study in hematopoietic stem-cell transplant recipients looked at the effects of antivirals on serum and cerebrospinal fluid HHV-6 viral load in patients diagnosed with encephalitis [85]. They found decrease in HHV-6 VL in both sites, concurrent with use of antiviral therapy (ganciclovir and/or foscarnet). However, this study did not provide perspective on possible differences in efficacy between the 2 agents [85]. In light of current available data, use of either agent appears appropriate for treatment of HHV-6 associated disease, including encephalitis. The AST IDCOP recommends treatment of HHV-6 associated encephalitis or other clinical syndrome attributed to HHV-6 but treatment of asymptomatic infections of the organ transplant host is not recommended [86]. Data on impact of current post-transplant antiviral prophylaxis on non-CMV viruses is limited. Available data suggests that the incidence of HHV-6, HHV-8, Varicella zoster virus (VZV) and Epstein-Barr virus (EBV) may be reduced but HHV-7 may not be impacted [87] Since most HHV-6 infections are asymptomatic and self-limited, and antiviral prophylaxis for HHV-6 following organ transplantation is currently not recommended [86]. Limited data suggests HHV-6 viremia threshold >104 copies/ml is predictive of encephalitis in hematopoietic stem-cell transplant recipients, however similar data is lacking in SOT [88]. If data were to emerge on the impact of asymptomatic HHV-6 infection in the organ transplant host, the recommendation for antiviral prophylaxis and/or preemptive treatment of asymptomatic infection may change. However, any such approach would need to take into account the risk from toxicity of currently available antiviral agents.

The emergence of ganciclovir-resistant HHV-6 strains during treatment has been reported which correlates with U69 and U38 mutations [89,90]. Cidofovir resistance in HHV-6 has also been shown through in-vitro selection and is associated with mutation in U38 [91]. The emergence of drug-resistance HHV-6 strain should be considered if indicated clinically, during antiviral prophylaxis or treatment for other herpes viruses such as CMV. Further study is however, required to better understand the clinical significance of these drug resistant strains.

4. Human Herpesvirus-7

HHV-7 is a member of the herpesvirinae family and β -herpesvirus sub-family. It was discovered in 1990 from human CD4 T-cells [92]. HHV-7 is a ubiquitous virus that exhibits similar cellular tropism to HHV-6 viruses but appears to be less pathogenic. Similar to HHV-6B, over 90% of the adult population is estimated to be seropositive. Infection due to HHV-7 occurs in early childhood but tends to occur later than HHV-6B [93]. The clinical manifestations of primary HHV-7 infection are not as well-defined but it has been described as a cause of exanthem subitum and febrile illness with seizures [94,95]. Primary HHV-7 infection is followed by the establishment life-long latency in host-peripheral blood mononuclear cells [96].

4.1 Clustering of Beta-Herpesviruses - Co-Pathogens with CMV or ‘Innocent Bystanders’

Reactivation of HHV-7 following organ transplantation has been well-documented however, as is the case with HHV-6, search for direct pathogenicity has been challenging due to overlapping clinical manifestations, clustering  of viral reactivation with CMV, and correlation with occurrence of CMV disease [97]. Many authors have described HHV-6 and 7 as co-pathogens with CMV. Interaction between the 3 β-herpesviruses has been postulated to play a role in both the direct and indirect viral effects attributed to CMV [52]. Contrary to this, more recent prospective data while showing high prevalence of herpesvirus co-infection in SOT recipients with CMV disease, failed to show significant impact of HHV-6/7 co-infection [98]. In this study by Human et al, response to CMV antiviral therapy, baseline CMV viral load, time to viral eradication and risk of recurrence in patients with HHV-6 or7 co-infection and in those without, were similar [98]. The findings of this study put into question the clinical impact of HHV-6/7 co-infections on CMV infection/disease in organ transplant recipients and hence despite high prevalence, routine post-transplant monitoring for these viruses in not recommended [86]. There continue to be no reliable reports of clinical syndromes solely attributable to HHV-7 in organ transplant recipients.

4.2 Impact of Routine Antiviral Prophylaxis on HHV-7

Antiviral prophylaxis with oral ganciclovir or valganciclovir does not appear to have an impact on HHV-7 [87,98]. In-vitro measurement of end-point inhibitory concentrations (EC-50) for antivirals against β-herpesviruses revealed that HHV-7 was most susceptible to cidofovir but the EC-50 was higher than for HHV-6A/B (0.3 µg/ml for HHV-6A, 1.2 µg/ml for HHV-6B and 3.0 µg/ml for HHV-7) [99]. The clinical relevance of this data is unclear and there are no clinical interpretative breakpoints for such antiviral susceptibility testing.

5. Human Herpesvirus-8

Human herpesvirus-8 (HHV-8) is a gamma-herpesvirus [100]. As with all herpesviruses, HHV-8 establishes a lifelong state of latency following initial infection, and in the presence of certain host factors can enter the lytic phase producing actively infective virions which can kill the host cell. HHV-8 has a broad range of cellular tropism and natural reservoir for infection includes lymphoid, epithelial and endothelial cells [100].

HHV-8 seroprevelance in the world varies greatly by geographic region, with the highest rates of infection found in the Africa (up to 58%), Mediterranean (14%-25%), and the Middle East [101,102,103,104,105,106,107].

In endemic areas infection is acquired in childhood, in comparison to areas of low seroprevelance where it appears to be acquired later in life [102,108]. HHV-8 transmission occurs through saliva, blood, sexual contact and organ transplantation [109,110,111]. In non-endemic areas, high seroprevelance has been found among men who have sex with men (MSM) [112,113,114], and migrants from endemic areas [115]. Global seroprevelance of HHV-8 among MSM has been estimated in a recent meta-analysis at 33.0% (95%CI 29.2%-37.1%) [116]. In the setting of organ transplantation, infection can develop due to secondary reactivation in a seropositive recipient but primary HHV-8 infection due to donor and non-donor derived exposures has been well-described [110,111].

5.1 Clinical Manifestations of HHV-8: Oncogenic and Non-Oncogenic Properties

HHV-8 remains best known as an oncogenic virus and the causal agent of Kaposi’s sarcoma (KS), multi-centric Castleman’s disease (MCD) and primary effusion lymphoma (PEL).

Pre-existing HHV-8 seropositivity in the transplant population in endemic areas has been reported in the range of 4.9% to 16% [117,118,119]. The frequency of KS in solid-organ transplant recipients parallels the seroprevelance of HHV-8 infection in different regions of the world. In areas where HHV-8 infection is endemic, KS is a commonly seen malignancy in the transplant host; in one report from Saudi Arabia accounting for nearly 88% of all post-transplant cancers [120]. In a more recent report from Turkey during 1997 to 2017, KS was identified as the second most common post-transplant malignancy accounting for 18% of the cases, following squamous and basal cell carcinoma which were the most common at 29% [121]. The overall KS risk following solid-organ transplantation has been shown to be 125.3-fold higher (95% CI, 98.2–157.6) than in the general Italian population, reflecting the risk conferred by HHV-8 infection in endemic areas [122]. In contrast, in areas of low seroprevelance such as the United States, the prevalence of post-transplant KS is much lower; and the most common malignancies in solid-organ transplant recipients are non-Hodgkin lymphoma, lung, liver and kidney cancer [123]. Analysis of United States solid-organ transplant and population based cancer registries, identified 163 cases of KS among 264,624 transplant recipients from 1987-2014; with incidence rate of 12.4 per 100,000 person-years [124]. The same study also showed a decreasing incidence rate of KS with passage of time post-transplant (<1 year post-transplant: 35.1, <3: 15.6, <5: 5.0 and ≥ 5: 2.6 per 100, 000 person years); and a decrease in incidence rate during more recent years (1997-2003: 13.8 and 2004-2014: 9.6 per 100,000 person years) [124].

HHV-8 infection at the time of transplant is the most important risk factor for post-transplant KS [119,122,125,126]. There appears to be a difference in the KS risk conferred by preexisting/pre-transplant versus acquired/post-transplant HHV-8 infection; with higher risk in patients with pre-transplant HHV-8 infection [119,126]. The rates of post-transplant KS among HHV-8 seropositive kidney transplant recipients and seronegative recipients with positive donors (D+/R-), are 13% and 4.6% respectively [126]. These findings suggests that reactivation of latent HHV-8 infection plays a greater role in the development of post-transplant KS than incident infection. Other risk factors identified in majority of studies include increased recipient age at transplant, male sex, country of birth and ethnicity [127]. The role of induction and maintenance immunosuppressives with risk of KS remains unclear [127]. Some studies have shown a significant association with total amount of calcineurin inhibitor and use of anti-thymocyte globulin or IL-2 receptor antibody induction [127]; however the same has not been seen in other studies [124,127,128]. Similar to previous data, a more recent study by Cahoon et al showed increased risk of KS among males, non-white race, non-US citizenship, lung transplantation (versus kidney) and older age at the time of transplant (>65 years old = 28.6, 50-64 = 16.4, 35-49=7.7 and 15-34= 5.7 per 100, 000 person-years); and no association with specific induction or maintenance immunosuppressive medications [124].

KS is the most frequent manifestation of HHV-8 infection in the transplant host which presents in 90% of the cases as mucocutaneous lesions. According to Israel Penn International Transplant Tumor Registry (IPITTR) data, in 60% disease was limited to the skin, oropharyngeal mucosa and conjunctiva. Visceral disease has been reported in about 40% of cases with concomitant cutaneous KS in majority of patients [129,130]. More recently published OPTN data on US transplant recipients (1987-2003) identified 65 cases of KS, 80% of which were cutaneous-only and 20% with reported visceral disease [131]. Further analysis of the IPTTIR (1982 to 2001) was carried out to determine if there are clinical differences between post-transplant KS among international and US cases. This analysis revealed a higher rate of visceral KS among international patients in comparison to US patients who presented primarily with cutaneous-only disease (47% versus 27%, P <0.01) [132]. Among international patients, median time to presentation with post-transplant KS was shorter than among US patients and visceral KS was associated with significantly reduced 1-year patient survival in comparison with cutaneous-only disease (90% versus 70%, P <0.01). Therapeutic approach was similar and in both groups most commonly consisted of minimization of immunosuppression with either surgery and/or radiation therapy [132]. Limited data suggests that the clinical manifestations and treatment outcomes SOT recipients with KS is similar between those who have HIV infection and those without [133].

Less common oncologic manifestations of HHV-8 infection include multi-centric Castleman’s disease (MCD) and primary effusion lymphoma (PEL) [86,134,135]. Common clinical manifestations of MCD include fevers, pancytopenia, lymphadenopathy, elevated inflammatory markers and splenomegaly [136]. The diagnosis of PEL should be considered in a patient presenting with effusions in relation to serosal surfaces of the body (pericardium, pleural, peritoneal) [137,138]. Concomitant occurrence of multiple disease manifestations of HHV-8 has also been reported in transplant recipients including KS and MCD [139,140,141], and KS, MCD and HLH [142]. While HHV-8 remains best known as an oncogenic virus, there is increasing recognition of non-neoplastic disease manifestations including severe bone marrow suppression, multi-organ failure and hemophagocytic syndrome (HLH) [142,143,144,145,146,147].

5.2 Laboratory Testing for HHV-8: Role of Viral Testing in Screening, Diagnosis and Prognosis of HHV-8-Associated Diseases 

Most cells in HHV-8 associated disease are latently infected however, a proportion of cells do enter lytic replication (highest in MCD and lowest in KS); and both latent and lytic proteins play a role in tumorigenesis and inflammation [148,149]. Current diagnostics are based on detection of latent and/or lytic gene products through molecular or serologic methodology. In addition, immunohistochemical staining on tissue is available for diagnostic confirmation of HHV-8 associated disease. Below we outline the potential indications for HHV-8 serology and PCR in the organ transplant population.

5.2.1 Organ Donor and Recipient Screening

A number of commercial assays are available for HHV-8 serologic testing. These can be used to screen for latent HHV-8 infection in organ donors and recipients, but are not diagnostic of active infection. Certainly, such an approach appears attractive especially in HHV-8 endemic regions of the world, and could be utilized to risk stratify donors and recipients prior to transplantation. This approach is in fact, utilized by some transplant centers in endemic regions however; wider implementation of this practice continues to be impacted by lack of standardization in testing methodology and inter-laboratory variability in results. The continued lack of a gold-standard HHV-8 serologic assay was shown nicely in a recent multi-center prospective study which found, that out of 6 commercial serologic assays (both LANA and lytic antigen based), only 2 assays (both based on lytic antigen) showed perfect agreement with the reference standard [150]. In this study, the investigators also looked for baseline (pre-transplant) HHV-8 viremia using peripheral whole blood or plasma PCR testing. The correlation between baseline PCR and serologic test results was poor, both for donors and recipients; only 3 out 10 seropositive donors had detectable viremia at baseline, and only 1 out of 93 among seropositive recipients [150].

5.2.2 Diagnosis of HHV-8 Associated Disease (KS/PEL/MCD)

When HHV-8 associated disease is suspected, every effort must be made to obtain tissue specimens for diagnosis. Viral testing of tissue specimens using PCR and/or immunohistochemistry appears to carry the greatest diagnostic accuracy. The role of both peripheral blood PCR and serology in the diagnosis of HHV-8 associated disease remains controversial [149,151].

HHV-8 replication precedes the development of KS but the level of viremia is typically low. Blood PCR testing for diagnosis of KS therefore, in most studies has shown poor sensitivity and specificity [126,149,150,152,153]. Data on HIV/AIDS patients has shown that only ~ 10%-60% of patients with KS will have HHV-8 viremia [148,154,155]. Specific to SOT, data are limited but similar. A prospective investigation of HHV-8 associated complications in kidney transplant recipients reported that PCR had a sensitivity of only 23.8% in HHV-8 seropositive recipients. Among the 3 patients in this study who developed HHV-8 associated disease (2 KS and 1 non-neoplastic primary infection), but significant viremia was seen only in 1 patient supporting the low diagnostic yield of blood HHV-8 PCR in patients with KS [126].

Blood levels for HHV-8 are higher in PEL and MCD, and peripheral blood testing for viremia may have greater yield in diagnosis. In the absence of histopathology, HHV-8 viremia can be supportive however; histopathology remains the gold standard for diagnosis [148]. A recent study compared the degree of HHV-8 viremia between KS, PEL and MCD [156]. Similar to results from prior studies, patients with KS had the lowest levels of viremia. Among patients with KS, higher level viremia was seen in patients who had advanced stage KS and clinical progression of disease. Blood levels for HHV-8 were higher for patients with PEL and MCD and overall, the highest level of detectable virus was seen in effusion fluid in patients with PEL [156]. Based on these results, the authors postulated that there might be a role for HHV-8 viral load monitoring in patients with KS who are suspected of developing concomitant PEL and/or MCD [156].

5.2.3 Viral Load Monitoring

Detection of HHV-8 viremia in the blood however, has been shown to be helpful in predicting the risk of incident KS by providing a quantitative measure of lytic replication [154,155]; and has shown correlation with disease progression and regression in patients on treatment for KS [157]. Pellet et al reported results of quantitative HHV-8 PCR testing in 43 cases of post-transplant KS. In their cohort, 40% of patients were found to have significant HHV-8 viremia (> 100 copies per microgram of DNA) [157]. The authors conducted an analysis of factors associated with viremia and identified progressive disease due to KS as an independent predictor of viremia, suggesting a role of HHV-8 viral load testing as a prognostic marker in patients with post-transplant KS [157].

Available data on HHV-8 diagnostics must be reviewed bearing in mind that current testing modalities are not standardized. There continues to be significant variability in sensitivity and specificity between different HHV-8 blood assays and the clinical application of these in diagnosis of HHV-8 associated diseases is not clear in the absence of histopathology. Based on published literature, we make the following recommendations in regards to current state of HHV-8 diagnostics.

  • Peripheral blood HHV-8 serologic testing can be utilized to screen for asymptomatic infection and risk stratification in the transplant setting; however its utility in the diagnosis of HHV-8 associated disease is unclear.
  • Peripheral blood PCR for HHV-8 viremia appears to have greater yield in PEL/MCD and non-neoplastic HHV-8 related disease due to higher viral load in these conditions. Its role in the diagnosis of KS is limited due to typically low viral loads.
  • The current literature suggests that detection of HHV-8 viremia by blood PCR is most useful in predicting risk of incident disease and prognosis of patients with known HHV-8 associated diseases.
  • Diagnosis of HHV-8 associated disease must rely on histopathology with use of molecular and immunohistochemical testing for detection of HHV-8.

5.3 Monitoring and Treatment of HHV-8 Associated Diseases in Transplant Recipients

In the absence of robust data in SOT, current guidance regarding management of post-transplant HHV-8 associated diseases is based largely on expert opinion [86,149,158]. Riva et al proposed a comprehensive algorithm for monitoring and treatment of HHV-8 associated complications in the transplant recipient. They propose pre-transplant serologic screening of donors and recipients for stratification of risk, followed by monitoring of high risk recipients (D+/R- and R+) for viremia, virus-specific cell mediated immunity and clinical manifestations of HHV-8 associated disease [149]. Currently, donor and recipient HHV-8 serologic screening is performed in some centers mostly in endemic areas [149,159,160]. While such a practice seems useful as a tool for risk stratification and post-transplant surveillance, the lack of standardized testing has prevented this from being implemented on a larger scale. Nevertheless, recognition of HHV-8 risk in both the donor and recipient is essential for monitoring and timely recognition of related disease. Transplant practioners must therefore, obtain detailed history of risk factors for infection and perform targeted screening in individuals at high risk, such as those from regions endemic for HHV-8. In the event that clinical signs and symptoms of HHV-8 associated disease complications do occur, confirmation of diagnosis must be made by histopathology and a multi-disciplinary management approach including consultation with oncology should be pursued.

For all post-transplant HHV-8 associated, neoplastic and non-neoplastic diseases, the mainstay of treatment is reduction of immunosuppression and change to an mTOR inhibitor based regimen (sirolimus or everolimus) [149,158]. Study of patients with post-transplant KS has shown that the T-cell mediated immune response to HHV-8 is nearly absent at the onset of disease, with clinical regression occurring with reconstitution of virus-specific T-cell mediated immunity [161]. Over a decade now, data has been accumulating on the benefits of mTOR inhibitors in the management of post-transplant HHV-8 associated diseases [162,163,164,165]. Most case reports and series pertain to sirolimus; however successful management of post-transplant KS with switch to an everolimus based regimen has also been reported [162,163]. Sirolimus serves as a therapeutic agent in post-transplant KS due to direct anti-tumor activity (through inhibition of the mTOR pathway) and anti-angiogenetic activity (through inhibition of vascular endothelial growth factor) [166,167]. Rapamycin also has antiviral properties and is able to significantly inhibit HHV-8 lytic replication, providing further insight into the clinical benefit of these agents in patients with KS [168]. Reduction in immunosuppression alone is successful in many cases of post-transplant KS however, patients must be carefully followed for allograft rejection and need for chemotherapy (especially in cases of advanced disease) [158].

It is well-established that viral replication plays a key role in the pathogenesis of HHV-8 associated diseases. A number of antiviral agents have been found to have in-vitro activity against HHV-8 including, herpesvirus DNA synthesis inhibitors ganciclovir, valganciclovir, foscarnet and cidofovir [148,169]. However, a direct role of antiviral agents in the prevention and treatment of HHV-8 associated diseases is yet to be defined. Among antiviral agents with HHV-8 activity, valganciclovir is the only one that has been studied in a clinical trial [170]. In this study, the investigators studied the impact of valganciclovir on viral replication by measuring HHV-8 viral load in oropharyngeal specimens, in a cohort of 26 HHV-8 seropositive men. Of these, 16 were also HIV infected. In the valganciclovir arm, significant reduction in HHV-8 viral load was detected in comparison to the placebo arm (44% in placebo group versus 23% in valganciclovir group- RR 0.54 [95% CI 0.33–0.90]; P 0.02), establishing the efficacy of valganciclovir in reducing HHV-8 viral replication. Based on the findings of this study, one can postulate that suppression of HHV-8 viral replication using antiviral agents may have a role in prevention and/or treatment of KS, however larger scale clinical trials are needed. Beyond this study, data on these agents is limited to small observational studies which reports benefits of antiviral therapy in prevention and regression of KS [148].

In addition to herpesvirus DNA synthesis inhibitors, there has been considerable interest in the role of HIV protease inhibitors which have been associated with reduction in incidence and improving outcomes in patients with HIV/AIDS associated KS [171]. Further research into the mechanism behind these observations has shown anti-tumor activity independent of HIV protease inhibition through a broad range of cellular pathways [172,173]. Among the protease inhibitors, nelfinavir appears to be most promising agent due to direct antiviral activity against HHV-8 in addition to anti-neoplastic properties [174,175].

Based on currently available data, it is safe to conclude that no firm recommendations can be made in regards to use of antiviral agents in the management of HHV-8 associated disease in post-transplant patients. Data is limited in general, and mostly pertain to HIV/AIDS patients. The demonstration of valganciclovir efficacy in reducing HHV-8 replication in a clinical trial, and the discovery of novel anti-tumor property of nelfinavir, is certainly exciting; and hopefully future research will pave the path towards a defined role of these agents in the management of HHV-8 associated disease. Till then, current standard of care consisting of reduction in immunosuppression, use of a rapamycin based regimen, and chemotherapy in consultation with oncology should be pursued.

6. Conclusion

HHV-6,-7,-8 and HTLV-1 each pose unique challenges to the transplant host. Diagnosis of true infection due to HHV-6 and -7 remains challenging. In particular, clinical scenarios where HHV-6 and HHV-7 are detected with other beta-herpesviruses such as CMV; it is hard to know whether these are innocent bystanders or co-pathogens. Additionally, CIHHV-6 is often misdiagnosed as active HHV-6 infection. Education of transplant practioners on CIHHV-6, can help avoid unnecessary use of antivirals in patients. HHV-8 and HTLV-1 are well-recognized pathogens in the transplant host, but the uneven global distribution of these contributes to failed recognition of infection and associated complications; especially in non-endemic regions of the world. Development of standardized laboratory assays for HHV-8 and HTLV-1 is needed for accurate diagnosis and targeted screening of high risk organ donors and recipients.

Author Contributions

K.O. wrote the manuscript. S.T. reviewed, wrote, revised and formatted the manuscript.

Competing Interests

The authors declare no competing interests in regards to this manuscript.

References

  1. Gallo RC, Willems L, Hasegawa H. Screening transplant donors for HTLV-1 and -2. Blood. 2016; 128: 3029-3031. [CrossRef]
  2. Poiesz BJ, Ruscetti FW, Gazdar AF, Bunn PA, Minna JD, Gallo RC. Detection and isolation of type C retrovirus particles from fresh and cultured lymphocytes of a patient with cutaneous T-cell lymphoma. Proc Natl Acad Sci U S A. 1980; 77: 7415-7419. [CrossRef]
  3. Armstrong MJ, Corbett C, Rowe IA, Taylor GP, Neuberger JM. HTLV-1 in solid-organ transplantation: Current challenges and future management strategies. Transplantation. 2012; 94: 1075-1084. [CrossRef]
  4. Kaul DR, Davis JA. Human T cell lymphotrophic virus 1/2 in solid organ transplantation. Am J Transplant. 2013; 13: 355-360. [CrossRef]
  5. Proietti FA, Carneiro-Proietti AB, Catalan-Soares BC, Murphy EL. Global epidemiology of HTLV-I infection and associated diseases. Oncogene. 2005; 24: 6058-6068. [CrossRef]
  6. Mueller N, Okayama A, Stuver S, Tachibana N. Findings from the Miyazaki Cohort Study. J Acquir Immune Defic Syndr Hum Retrovirol.1996; 13: S2-S7. [CrossRef]
  7. Glynn SA, Kleinman SH, Schreiber GB, Busch MP, Wright DJ, Smith JW, et al. Trends in incidence and prevalence of major transfusion-transmissible viral infections in US blood donors, 1991 to 1996. REDS. 2000; 284: 229-235. [CrossRef]
  8. Hino S, Yamaguchi K, Katamine S, Sugiyama H, Amagasaki T, Kinoshita K, et al. Mother-to-child transmission of human T-cell leukemia virus type-I. Jpn J Cancer Res. 1985; 76: 474-480.
  9. Roucoux DF, Wang B, Smith D, Nass CC, Smith J, Hutching ST, et al. A prospective study of sexual transmission of human T lymphotropic virus (HTLV)-I and HTLV-II. J Infect Dis. 2005; 191: 1490-1497. [CrossRef]
  10. Martín-Dávila P, Fortún J, López-Vélez R, Norman F, Montes de Oca M, Zamarrón P, et al. Transmission of tropical and geographically restricted infections during solid-organ transplantation. Clin Microbiol Rev. 2008; 21: 60-96. [CrossRef]
  11. Montesdeoca Andrade MJ, Correa Diaz EP, Buestan ME. HTLV-1-associated myelopathy in a solid organ transplant recipient. BMJ Case Rep. 2016; 2016. pii: bcr2016215243. [CrossRef]
  12. Zarranz Imirizaldu JJ, Gomez Esteban JC, Rouco Axpe I, Perez Concha T, Velasco Juanes F, Allue Susaeta I, et al. Post-transplantation HTLV-1 myelopathy in three recipients from a single donor. J Neurol Neurosurg Psychiatr. 2003; 74: 1080-1084. [CrossRef]
  13. Ramanan P, Deziel PJ, Norby SM, Yao JD, Garza I, Razonable RR. Donor-transmitted HTLV-1-associated myelopathy in a kidney transplant recipient--case report and literature review. Am J Transplant. 2014; 14: 2417-2421. [CrossRef]
  14. Torres JA, Taimur S. Postrenal transplant human T-Cell lymphotropic virus type I-associated myelopathy/tropical spastic paraparesis: A case report and review of the literature. Transplant Direct. 2015; 1: e3. [CrossRef]
  15. Nagamine Y, Hayashi T, Kato Y, Horiuchi Y, Tanahashi N. Human T lymphotropic virus type-1-associated myelopathy manifesting shortly after living-donor renal transplantation. Intern Med. 2015; 54: 75-78. [CrossRef]
  16. Nakatsuji Y, Sugai F, Watanabe S, Kaido M, Koguchi K, Abe K, et al. HTLV-I-associated myelopathy manifested after renal transplantation. J Neurol Sci. 2000; 177: 154-156. [CrossRef]
  17. Kaul DR, Taranto S, Alexander C, Covington S, Marvin M, Nowicki M, et al. Donor screening for human T-cell lymphotrophic virus 1/2: Changing paradigms for changing testing capacity. Am J Transplant. 2010; 10: 207-213. [CrossRef]
  18. Marvin MR, Brock GN, Kwarteng K, Nagubandi R, Ravindra KV, Eng M, et al. Increasing utilization of human T-cell lymphotropic virus (+) donors in liver transplantation: Is it safe? Transplantation. 2009; 87: 1180-1190. [CrossRef]
  19. Shames BD, D'Alessandro AM, Sollinger HW. Human T-cell lymphotrophic virus infection in organ donors: a need to reassess policy? Am J Transplant. 2002; 2: 658-663. [CrossRef]
  20. Hoshida Y, Li T, Dong Z, Tomita Y, Yamauchi A, Hanai J, et al. Lymphoproliferative disorders in renal transplant patients in Japan. Int J Cancer. 2001; 91: 869-875. [CrossRef]
  21. Taylor GP. Human T-lymphotropic virus type 1 infection and solid organ transplantation. Rev Med Virol. 2018; 28. doi: 10.1002/rmv. [CrossRef]
  22. Sabouri AH, Saito M, Lloyd AL, Vine AM, Witkover AW, Furukawa Y, et al. Polymorphism in the interleukin-10 promoter affects both provirus load and the risk of human T lymphotropic virus type I-associated myelopathy/tropical spastic paraparesis. J Infect Dis. 2004; 190: 1279-1285. [CrossRef]
  23. Treviño A, Lopez M, Vispo E, Aguilera A, Ramos JM, Benito R, et al. Development of tropical spastic paraparesis in human T-lymphotropic virus type 1 carriers is influenced by interleukin 28B gene polymorphisms. Clin Infect Dis. 2012; 55: e1-e4. [CrossRef]
  24. Anderson DW, Epstein JS, Lee TH, Lairmore MD, Saxinger C, Kalyanaraman VS, et al. Serological confirmation of human T-lymphotropic virus type I infection in healthy blood and plasma donors. Blood. 1989; 74: 2585-2591.
  25. Liu H, Shah M, Stramer SL, Chen W, Weiblen BJ, Murphy EL. Sensitivity and specificity of human T-lymphotropic virus (HTLV) types I and II polymerase chain reaction and several serologic assays in screening a population with a high prevalence of HTLV-II. Transfusion. 1999; 39: 1185-1193. [CrossRef]
  26. Demontis MA, Sadiq MT, Golz S, Taylor GP. HTLV-1 viral RNA is detected rarely in plasma of HTLV-1 infected subjects. J Med Virol. 2015; 87: 2130-2134. [CrossRef]
  27. Iwanaga M, Watanabe T, Utsunomiya A, Okayama A, Uchimaru K, Koh KR, et al. Human T-cell leukemia virus type I (HTLV-1) proviral load and disease progression in asymptomatic HTLV-1 carriers: A nationwide prospective study in Japan. Blood. 2010; 116: 1211-1219. [CrossRef]
  28. Takenouchi N, Yamano Y, Usuku K, Osame M, Izumo S. Usefulness of proviral load measurement for monitoring of disease activity in individual patients with human T-lymphotropic virus type I-associated myelopathy/tropical spastic paraparesis. J Neurovirol. 2003; 9: 29-35. [CrossRef]
  29. Huang RC, Fishman JA. Screening of deceased organ donors: No easy answers. Transplantation. 2011; 91: 146-149. [CrossRef]
  30. WHO. Virus diseases: Human T lymphotropic virus type I, HTLV-I. Wkly Epidemiol Rec. 1989; 64: 382.
  31. Matsushita S, Mitsuya H, Reitz MS, Broder S. Pharmacological inhibition of in vitro infectivity of human T lymphotropic virus type I. J Clin Invest. 1987; 80: 394-400. [CrossRef]
  32. Seegulam ME, Ratner L. Integrase inhibitors effective against human T-cell leukemia virus type 1. Antimicrob Agents Chemother. 2011; 55: 2011-2017. [CrossRef]
  33. Taylor GP, Goon P, Furukawa Y, Green H, Barfield A, Mosley A, et al. Zidovudine plus lamivudine in human T-lymphotropic virus type-I-associated myelopathy: A randomised trial. Retrovirology. 2006; 3: 63. [CrossRef]
  34. Trevino A, Parra P, Bar-Magen T, Garrido C, de Mendoza C, Soriano V. Antiviral effect of raltegravir on HTLV-1 carriers. J Antimicrob Chemother. 2012; 67: 218-221. [CrossRef]
  35. Nakamura T, Matsuo T, Fukuda T, Yamato S, Yamaguchi K, Kinoshita I, et al. Efficacy of prosultiamine treatment in patients with human T lymphotropic virus type I-associated myelopathy/tropical spastic paraparesis: Results from an open-label clinical trial. BMC Med. 2013; 11: 182. [CrossRef]
  36. Matsuo T, Miyata Y, Nakamura T, Satoh K, Sakai H. Prosultiamine for treatment of lower urinary tract dysfunction accompanied by human T-lymphotropic virus type 1-associated myelopathy/tropical spastic paraparesis. Int J Urol. 2018; 25: 54-60. [CrossRef]
  37. Nishiura Y, Nakamura T, Fukushima N, Nakamura H, Ida H, Aramaki T, et al. Disulfide-mediated apoptosis of human T-lymphotrophc virus type-I (HTLV-I)-infected cells in patients with HTLV-I-associated myelopathy/tropical spastic paraparesis. Antivir Ther. 2009; 14: 533-542.
  38. Salahuddin SZ, Ablashi DV, Markham PD, Josephs SF, Sturzenegger S, Kaplan M, et al. Isolation of a new virus, HBLV, in patients with lymphoproliferative disorders. Science. 1986; 234: 596-601. [CrossRef]
  39. Ablashi D, Agut H, Alvarez-Lafuente R, Clark DA, Dewhurst S, DiLuca D, et al. Classification of HHV-6A and HHV-6B as distinct viruses. Arch Virol. 2014; 159: 863-870. [CrossRef]
  40. Hall CB, Long CE, Schnabel KC, Caserta MT, McIntyre KM, Costanzo MA, et al. Human herpesvirus-6 infection in children. A prospective study of complications and reactivation. N Engl J Med. 1994; 331: 432-438. [CrossRef]
  41. Pruksananonda P, Hall CB, Insel RA, McIntyre K, Pellett PE, Long CE, et al. Primary human herpesvirus 6 infection in young children. N Engl J Med. 1992; 326: 1445-1450. [CrossRef]
  42. Zerr DM, Meier AS, Selke SS, Frenkel LM, Huang ML, Wald A, et al. A population-based study of primary human herpesvirus 6 infection. N Engl J Med. 2005; 352: 768-776. [CrossRef]
  43. Dewhurst S, McIntyre K, Schnabel K, Hall CB. Human herpesvirus 6 (HHV-6) variant B accounts for the majority of symptomatic primary HHV-6 infections in a population of U.S. infants. J Clin Microbiol. 1993; 31: 416-418.
  44. Agut H, Bonnafous P, Gautheret-Dejean A. Laboratory and clinical aspects of human herpesvirus 6 infections. Clin Microbiol Rev. 2015; 28: 313-335. [CrossRef]
  45. Flamand L. Pathogenesis from the reactivation of chromosomally integrated human herpesvirus type 6: Facts rather than fiction. Clin Infect Dis. 2014; 59: 549-551. [CrossRef]
  46. Leong HN, Tuke PW, Tedder RS, Khanom AB, Eglin RP, Atkinson CE, et al. The prevalence of chromosomally integrated human herpesvirus 6 genomes in the blood of UK blood donors. J Med Virol. 2007; 79: 45-51. [CrossRef]
  47. Pellett PE, Ablashi DV, Ambros PF, Agut H, Caserta MT, Descamps V, et al. Chromosomally integrated human herpesvirus 6: Questions and answers. Rev Med Virol. 2012; 22: 144-155. [CrossRef]
  48. Cervera C, Marcos MA, Linares L, Roig E, Benito N, Pumarola T, et al. A prospective survey of human herpesvirus-6 primary infection in solid organ transplant recipients. Transplantation. 2006; 82: 979-982. [CrossRef]
  49. Ohashi M, Sugata K, Ihira M, Asano Y, Egawa H, Takada Y, et al. Human herpesvirus 6 infection in adult living related liver transplant recipients. Liver Transplant. 2008; 14: 100-109. [CrossRef]
  50. Rossi C, Delforge ML, Jacobs F, Wissing M, Pradier O, Remmelink M, et al. Fatal primary infection due to human herpesvirus 6 variant A in a renal transplant recipient. Transplantation. 2001; 71: 288-292. [CrossRef]
  51. Pilmore H, Collins J, Dittmer I, Williams L, Carpenter L, Thomas S, et al. Fatal human herpesvirus-6 infection after renal transplantation. Transplantation. 2009; 88: 762-765. [CrossRef]
  52. Razonable RR, Paya CV. The impact of human herpesvirus-6 and -7 infection on the outcome of liver transplantation. Liver Transplant. 2002; 8: 651-658. [CrossRef]
  53. Lee SO, Brown RA, Razonable RR. Chromosomally integrated human herpesvirus-6 in transplant recipients. Transpl Infect Dis. 2012; 14: 346-354. [CrossRef]
  54. Humar A, Kumar D, Caliendo AM, Moussa G, Ashi-Sulaiman A, Levy G, et al. Clinical impact of human herpesvirus 6 infection after liver transplantation. Transplantation. 2002; 73: 599-604. [CrossRef]
  55. Lautenschlager I, Hockerstedt K, Linnavuori K, Taskinen E. Human herpesvirus-6 infection after liver transplantation. Clin Infect Dis. 1998; 26: 702-707. [CrossRef]
  56. Razonable RR, Rivero A, Brown RA, Hart GD, Espy MJ, van Cruijsen H, et al. Detection of simultaneous beta-herpesvirus infections in clinical syndromes due to defined cytomegalovirus infection. Clin Transplant. 2003; 17: 114-120. [CrossRef]
  57. Singh N, Carrigan DR, Gayowski T, Marino IR. Human herpesvirus-6 infection in liver transplant recipients: documentation of pathogenicity. Transplantation. 1997; 64: 674-678. [CrossRef]
  58. Potenza L, Luppi M, Barozzi P, Rossi G, Cocchi S, Codeluppi M, et al. HHV-6A in syncytial giant-cell hepatitis. N Engl J Med. 2008; 359: 593-602. [CrossRef]
  59. Guardia AC, Stucchi RS, Sampaio AM, Milan A, Costa SC, Pavan CR, et al. Human herpesvirus 6 in donor biopsies associated with the incidence of clinical cytomegalovirus disease and hepatitis C virus recurrence. Int J Infect Dis. 2012; 16: e124-129. [CrossRef]
  60. Singh N, Husain S, Carrigan DR, Knox KK, Weck KE, Wagener MM, et al. Impact of human herpesvirus-6 on the frequency and severity of recurrent hepatitis C virus hepatitis in liver transplant recipients. Clin Transplant. 2002; 16: 92-96. [CrossRef]
  61. Lautenschlager I, Lappalainen M, Linnavuori K, Suni J, Hockerstedt K. CMV infection is usually associated with concurrent HHV-6 and HHV-7 antigenemia in liver transplant patients. J Clin Virol. 2002; 25: S57-S61. [CrossRef]
  62. Sampaio AM, Guardia AC, Milan A, Sasaki AN, Andrade PD, Bonon SH, et al. Co-infection and clinical impact of human herpesvirus 5 and 6 in liver transplantation. Transplant Proc. 2012; 44: 2455-2458. [CrossRef]
  63. Harma M, Hockerstedt K, Lyytikainen O, Lautenschlager I. HHV-6 and HHV-7 antigenemia related to CMV infection after liver transplantation. J Med Virol. 2006; 78: 800-805. [CrossRef]
  64. Chapenko S, Folkmane I, Ziedina I, Chistyakovs M, Rozentals R, Krumina A, et al. Association of HHV-6 and HHV-7 reactivation with the development of chronic allograft nephropathy. J Clin Virol. 2009; 46: 29-32. [CrossRef]
  65. Rogers J, Rohal S, Carrigan DR, Kusne S, Knox KK, Gayowski T, et al. Human herpesvirus-6 in liver transplant recipients: Role in pathogenesis of fungal infections, neurologic complications, and outcome. Transplantation. 2000; 69: 2566-2573. [CrossRef]
  66. Neurohr C, Huppmann P, Leuchte H, Schwaiblmair M, Bittmann I, Jaeger G, et al. Human herpesvirus 6 in bronchalveolar lavage fluid after lung transplantation: a risk factor for bronchiolitis obliterans syndrome? Am J Transplant. 2005; 5: 2982-2991. [CrossRef]
  67. Lawrence GL, Chee M, Craxton MA, Gompels UA, Honess RW, Barrell BG. Human herpesvirus 6 is closely related to human cytomegalovirus. J Virol. 1990; 64: 287-299.
  68. Nasimfar A, Sadeghi E, Alborzi A, Sepehrvand N, Ziyaeyan M, Jamalidoust M, et al. The activation of cytomegalovirus and human herpes virus 6 after liver transplantation. Hepat Mon. 2018; 2: e11987. [CrossRef]
  69. Lautenschlager I, Krueger G, Ablashi. Human herpesviruses HHV-6A, HHV-6B & HHV-7. 3rd ed. Netherlands: Elsevier Science. p. 9-34
  70. Norton RA, Caserta MT, Hall CB, Schnabel K, Hocknell P, Dewhurst S. Detection of human herpesvirus 6 by reverse transcription-PCR. J Clin Microbiol. 1999; 37: 3672-3675.
  71. Loginov R, Karlsson T, Hockerstedt K, Ablashi D, Lautenschlager I. Quantitative HHV-6B antigenemia test for the monitoring of transplant patients. Eur J Clin Microbiol Infect Dis. 2010; 29: 881-886. [CrossRef]
  72. Achour A, Boutolleau D, Slim A, Agut H, Gautheret-Dejean A. Human herpesvirus-6 (HHV-6) DNA in plasma reflects the presence of infected blood cells rather than circulating viral particles. J Clin Virol. 2007; 38: 280-285. [CrossRef]
  73. Caserta MT, Hall CB, Schnabel K, Lofthus G, Marino A, Shelley L, et al. Diagnostic assays for active infection with human herpesvirus 6 (HHV-6). J Clin Virol. 2010; 48: 55-57. [CrossRef]
  74. Karlsson T, Mannonen L, Loginov R, Lappalainen M, Hockerstedt K, Lautenschlager I. Development of a new quantitative real-time HHV-6-PCR and monitoring of HHV-6 DNAaemia after liver transplantation. J Virol Methods. 2012; 181: 25-36. [CrossRef]
  75. Clark DA, Nacheva EP, Leong HN, Brazma D, Li YT, Tsao EH, et al. Transmission of integrated human herpesvirus 6 through stem cell transplantation: Implications for laboratory diagnosis. J Infect Dis. 2006; 193: 912-916. [CrossRef]
  76. Ward KN, Leong HN, Nacheva EP, Howard J, Atkinson CE, Davies NW, et al. Human herpesvirus 6 chromosomal integration in immunocompetent patients results in high levels of viral DNA in blood, sera, and hair follicles. J Clin Microbiol. 2006; 44: 1571-1574. [CrossRef]
  77. Potenza L, Barozzi P, Masetti M, Pecorari M, Bresciani P, Gautheret-Dejean A, et al. Prevalence of human herpesvirus-6 chromosomal integration (CIHHV-6) in Italian solid organ and allogeneic stem cell transplant patients. Am J Transplant. 2009; 9: 1690-1697. [CrossRef]
  78. Zhang E, Bell AJ, Wilkie GS, Suárez NM, Batini C, Veal CD, et al. Inherited chromosomally integrated human herpesvirus 6 genomes are ancient, intact, and potentially able to reactivate from telomeres. J Virol. 2017; 91: e01137-17 [CrossRef]
  79. Clark DA, Nacheva EP, Leong HN, Brazma D, Li YT, Tsao EH, et al. Transmission of integrated human herpesvirus 6 through stem cell transplantation: Implications for laboratory diagnosis. J Infect Dis. 2006; 193: 912-916. [CrossRef]
  80. De Bolle L, Naesens L, De Clercq E. Update on human herpesvirus 6 biology, clinical features, and therapy. Clin Microbiol Rev. 2005; 18: 217-245. [CrossRef]
  81. Lamoth F, Jayet PY, Aubert JD, Rotman S, Mottet C, Sahli R, et al. Case report: Human herpesvirus 6 reactivation associated with colitis in a lung transplant recipient. J Med Virol. 2008; 80: 1804-1807. [CrossRef]
  82. Nash PJ, Avery RK, Tang WH, Starling RC, Taege AJ, Yamani MH. Encephalitis owing to human herpesvirus-6 after cardiac transplant. Am J Transplant. 2004; 4: 1200-1203. [CrossRef]
  83. Ongradi J, Ablashi DV, Yoshikawa T, Stercz B, Ogata M. Roseolovirus-associated encephalitis in immunocompetent and immunocompromised individuals. J Neurovirol. 2017; 23: 1-19. [CrossRef]
  84. Vinnard C, Barton T, Jerud E, Blumberg E. A report of human herpesvirus 6-associated encephalitis in a solid organ transplant recipient and a review of previously published cases. Liver Transplant. 2009; 15: 1242-1246. [CrossRef]
  85. Zerr DM, Gupta D, Huang ML, Carter R, Corey L. Effect of antivirals on human herpesvirus 6 replication in hematopoietic stem cell transplant recipients. Clin Infect Dis. 2002; 34: 309-317. [CrossRef]
  86. Razonable RR. Human herpesviruses 6, 7 and 8 in solid organ transplant recipients. Am J Transplant. 2013; 13: 67-77; quiz 77-68. [CrossRef]
  87. Razonable RR, Brown RA, Humar A, Covington E, Alecock E, Paya CV. Herpesvirus infections in solid organ transplant patients at high risk of primary cytomegalovirus disease. J Infect Dis. 2005; 192: 1331-1339. [CrossRef]
  88. Ogata M, Satou T, Kawano R, Goto K, Ikewaki J, Kohno K, et al. Plasma HHV-6 viral load-guided preemptive therapy against HHV-6 encephalopathy after allogeneic stem cell transplantation: a prospective evaluation. Bone Marrow Transplant. 2008; 41: 279-285. [CrossRef]
  89. Isegawa Y, Hara J, Amo K, Osugi Y, Takemoto M, Yamanishi K, et al. Human herpesvirus 6 ganciclovir-resistant strain with amino acid substitutions associated with the death of an allogeneic stem cell transplant recipient. J Clin Virol. 2009; 44: 15-19. [CrossRef]
  90. Manichanh C, Olivier-Aubron C, Lagarde JP, Aubin JT, Bossi P, Gautheret-Dejean A, et al. Selection of the same mutation in the U69 protein kinase gene of human herpesvirus-6 after prolonged exposure to ganciclovir in vitro and in vivo. J Gen Virol. 2001; 82: 2767-2776. [CrossRef]
  91. Bonnafous P, Boutolleau D, Naesens L, Deback C, Gautheret-Dejean A, Agut H. Characterization of a cidofovir-resistant HHV-6 mutant obtained by in vitro selection. Antiviral Res. 2008; 77: 237-240. [CrossRef]
  92. Frenkel N, Schirmer EC, Wyatt LS, Katsafanas G, Roffman E, Danovich RM, et al. Isolation of a new herpesvirus from human CD4+ T cells. Proc Natl Acad Sci USA. 1990; 87: 748-752. [CrossRef]
  93. Agut H, Bonnafous P, Gautheret-Dejean A. Update on infections with human herpesviruses 6A, 6B, and 7. Med Mal Infect. 2017; 47: 83-91. [CrossRef]
  94. Ward KN, Andrews NJ, Verity CM, Miller E, Ross EM. Human herpesviruses-6 and -7 each cause significant neurological morbidity in Britain and Ireland. Arch Dis Child. 2005; 90: 619-623. [CrossRef]
  95. Freitas RB, Freitas MR, Oliveira CS, Linhares AC. Human herpesvirus-7 as a cause of exanthematous ilnesses in Belem, Para, Brazil. Rev Inst Med Trop Sao Paulo. 2004; 46: 139-143. [CrossRef]
  96. Katsafanas GC, Schirmer EC, Wyatt LS, Frenkel N. In vitro activation of human herpesviruses 6 and 7 from latency. Proc Natl Acad Sci USA. 1996; 93: 9788-9792. [CrossRef]
  97. Mendez JC, Dockrell DH, Espy MJ, Smith TF, Wilson JA, Harmsen WS, et al. Human beta-herpesvirus interactions in solid organ transplant recipients. J Infect Dis. 2001; 183: 179-184. [CrossRef]
  98. Humar A, Asberg A, Kumar D, Hartmann A, Moussa G, Jardine A, et al. An assessment of herpesvirus co-infections in patients with CMV disease: Correlation with clinical and virologic outcomes. Am J Transplant. 2009; 9: 374-381. [CrossRef]
  99. Yoshida M, Yamada M, Tsukazaki T, Chatterjee S, Lakeman FD, Nii S, Whitley RJ. Comparison of antiviral compounds against human herpesvirus 6 and 7. Antivir Res. 1998; 40: 73-84. [CrossRef]
  100. Edelman DC. Human herpesvirus 8--a novel human pathogen. Antivir Res. 2005; 2: 78.
  101. Biryahwaho B, Dollard SC, Pfeiffer RM, Shebl FM, Munuo S, Amin MM, et al. Sex and geographic patterns of human herpesvirus 8 infection in a nationally representative population-based sample in Uganda. J Infect Dis. 2010; 202: 1347-1353. [CrossRef]
  102. Butler LM, Were WA, Balinandi S, Downing R, Dollard S, Neilands TB, et al. Human herpesvirus 8 infection in children and adults in a population-based study in rural Uganda. J Infect Dis. 2011; 203: 625-634. [CrossRef]
  103. Rohner E, Wyss N, Trelle S, Mbulaiteye SM, Egger M, Novak U, et al. HHV-8 seroprevalence: A global view. Syst Rev. 2014; 3: 11. [CrossRef]
  104. Whitby D, Luppi M, Barozzi P, Boshoff C, Weiss RA, Torelli G. Human herpesvirus 8 seroprevalence in blood donors and lymphoma patients from different regions of Italy. J Natl Cancer Inst. 1998; 90: 395-397. [CrossRef]
  105. Cattani P, Cerimele F, Porta D, Graffeo R, Ranno S, Marchetti S, et al. Age-specific seroprevalence of human herpesvirus 8 in Mediterranean regions. Clin Microbiol Infect. 2003; 9: 274-279. [CrossRef]
  106. Dollard SC, Butler LM, Jones AM, Mermin JH, Chidzonga M, Chipato T, et al. Substantial regional differences in human herpesvirus 8 seroprevalence in sub-Saharan Africa: Insights on the origin of the "Kaposi's sarcoma belt". Int J Cancer. 2010; 127: 2395-2401. [CrossRef]
  107. Mbulaiteye SM, Pfeiffer RM, Whitby D, Brubaker GR, Shao J, Biggar RJ. Human herpesvirus 8 infection within families in rural Tanzania. J Infect Dis. 2003; 187: 1780-1785. [CrossRef]
  108. Blauvelt A, Sei S, Cook PM, Schulz TF, Jeang KT. Human herpesvirus 8 infection occurs following adolescence in the United States. J Infect Dis. 1997; 176: 771-774. [CrossRef]
  109. Crabtree KL, Wojcicki JM, Minhas V, Kankasa C, Mitchell C, Wood C. Association of household food- and drink-sharing practices with human herpesvirus 8 seroconversion in a cohort of zambian children. J Infect Dis. 2017; 216: 842-849. [CrossRef]
  110. Jenkins FJ, Hoffman LJ, Liegey-Dougall A. Reactivation of and primary infection with human herpesvirus 8 among solid-organ transplant recipients. J Infect Dis. 2002; 185: 1238-1243. [CrossRef]
  111. Regamey N, Tamm M, Wernli M, Witschi A, Thiel G, Cathomas G, et al. Transmission of human herpesvirus 8 infection from renal-transplant donors to recipients. New Engl J Med. 1998; 339: 1358-1363. [CrossRef]
  112. Regamey N, Cathomas G, Schwager M, Wernli M, Harr T, Erb P. High human herpesvirus 8 seroprevalence in the homosexual population in Switzerland. N Engl J Med. 1998; 36: 1784-1786. [CrossRef]
  113. Engels EA, Atkinson JO, Graubard BI, McQuillan GM, Gamache C, Mbisa G, et al. Risk factors for human herpesvirus 8 infection among adults in the United States and evidence for sexual transmission. J Infect Dis. 2007; 196: 199-207. [CrossRef]
  114. Dukers NH, Renwick N, Prins M, Geskus RB, Schulz TF, Weverling GJ, et al. Risk factors for human herpesvirus 8 seropositivity and seroconversion in a cohort of homosexual men. Am J Epidemiol. 2000; 151: 213-224. [CrossRef]
  115. de Tejada BM, Steffen I, Cantero P, Posfay-Barbe KM, Irion O, Hirschel B, et al. Human herpes virus type 8 seroprevalence in pregnant women in Geneva, Switzerland. J Matern Fetal Neonatal Med. 2011; 24: 183-185. [CrossRef]
  116. Liu Z, Fang Q, Zuo J, Chen Y, Minhas V, Wood C, Zhang T. Global epidemiology of human herpesvirus 8 in men who have sex with men: A systematic review and meta-analysis. J Med Virol. 2018; 90: 582-591. [CrossRef]
  117. Diociaiuti A, Nanni G, Cattani P, Lesnoni La Parola I, Masini MC, Capuano M, et al. HHV8 in renal transplant recipients. Transplant Int. 2000; 13: S410-S412. [CrossRef]
  118. Bergallo M, Costa C, Margio S, Sidoti F, Re D, Segoloni GP, Cavallo R. Human herpes virus 8 infection in kidney transplant patients from an area of northwestern Italy (Piemonte region). Nephrol Dial Transplant. 2007; 22: 1757-1761. [CrossRef]
  119. Andreoni M, Goletti D, Pezzotti P, Pozzetto A, Monini P, Sarmati L, et al. Prevalence, incidence and correlates of HHV-8/KSHV infection and Kaposi's sarcoma in renal and liver transplant recipients. J Infect. 2001; 43: 195-199. [CrossRef]
  120. Qunibi W, Akhtar M, Sheth K, Ginn HE, Al-Furayh O, DeVol EB, Taher S. Kaposi's sarcoma: the most common tumor after renal transplantation in Saudi Arabia. Am J Med. 1988; 84: 225-232. [CrossRef]
  121. Tuncer Vural A, Karatas Togral A, Gulec AT, Haberal M. Kaposi Sarcoma in the era of rapamycin remains a therapeutic challenge in organ transplant recipients. Exp Clin Transplant. 2018; 16: 25-27.
  122. Piselli P, Busnach G, Citterio F, Frigerio M, Arbustini E, Burra P, et al. Risk of Kaposi sarcoma after solid-organ transplantation: Multicenter study in 4,767 recipients in Italy, 1970-2006. Transplant Proc. 2009; 41: 1227-1230. [CrossRef]
  123. Engels EA, Pfeiffer RM, Fraumeni JF Jr, Kasiske BL, Israni AK, Snyder JJ, et al. Spectrum of cancer risk among US solid organ transplant recipients. JAMA. 2011; 306: 1891-1901. [CrossRef]
  124. Cahoon EK, Linet MS, Clarke CA, Pawlish KS, Engels EA, Pfeiffer RM. Risk of Kaposi sarcoma after solid organ transplantation in the United States. Int J Cancer. 2018; 143: 2741-2748. [CrossRef]
  125. Farge D, Lebbé C, Marjanovic Z, Tuppin P, Mouquet C, Peraldi MN, et al. Human herpes virus-8 and other risk factors for Kaposi's sarcoma in kidney transplant recipients. Groupe cooperatif de transplantation d' Ile de France (GCIF). Transplantation. 1999; 67: 1236-1242. [CrossRef]
  126. Francès C, Marcelin AG, Legendre Ch, Chevret S, Dussaix E, Lejeune J, et al. The impact of preexisting or acquired Kaposi sarcoma herpesvirus infection in kidney transplant recipients on morbidity and survival. Am J Transplant. 2009; 9: 2580-2586. [CrossRef]
  127. Hosseini-Moghaddam SM, Soleimanirahbar A, Mazzulli T, Rotstein C, Husain S. Post renal transplantation Kaposi's sarcoma: A review of its epidemiology, pathogenesis, diagnosis, clinical aspects, and therapy. Transpl Infect Dis. 2012; 14: 338-345. [CrossRef]
  128. Alzahrani AJ, El-Harith el-HA, Milzer J, Obeid OE, Stuhrmann M, Al-Dayel A, et al. Increased seroprevalence of human herpes virus-8 in renal transplant recipients in Saudi Arabia. Nephrol Dial Transplant. 2005; 20: 2532-2536. [CrossRef]
  129. Penn I. Kaposi's sarcoma in transplant recipients. Transplantation. 1997; 64: 669-673. [CrossRef]
  130. Penn I. Sarcomas in organ allograft recipients. Transplantation. 1995; 60: 1485-1491. [CrossRef]
  131. Mbulaiteye SM, Engels EA. Kaposi's sarcoma risk among transplant recipients in the United States (1993-2003). Int J Cancer. 2006; 119: 2685-2691. [CrossRef]
  132. Woodle ES, Hanaway M, Buell J, Gross T, First MR, Trofe J, et al. Kaposi sarcoma: An analysis of the US and international experiences from the Israel Penn International Transplant Tumor Registry. Transplant Proc. 2001; 33: 3660-3661. [CrossRef]
  133. Charpentier C, Delyon J, Glotz D, Peraldi MN, Rerolle JP, Barrou B, et al. Kaposi sarcoma in HIV-positive solid-organ transplant recipients: A french multicentric national study and literature review. Transplantation. 2019; 103: e22-e28. [CrossRef]
  134. Le J. Oncogenic gamma herpesviruses EBV and HHV8 in kidney transplantation. Semin Dialysis. 2016; 36: 362-371. [CrossRef]
  135. Matsunami M, Ubara Y, Sumida K, Oshima Y, Oguro M, Kinoshita K, et al. The efficacy and safety of anti-interleukin-6 receptor monoclonal blockade in a renal transplant patient with Castleman disease: Early post-transplant outcome. BMC Nephrol. 2018; 19: 263. [CrossRef]
  136. Bonatti HJ, Axt J, Hunter EB, Lott SL, Frangoul H, Gillis L, et al. Castleman disease in a pediatric liver transplant recipient: A case report and literature review. Pediatr Transplant. 2012; 16: E229-E234. [CrossRef]
  137. Kugasia IAR, Kumar A, Khatri A, Saeed F, Islam H, Epelbaum O. Primary effusion lymphoma of the pleural space: Report of a rare complication of cardiac transplant with review of the literature. Transpl Infect Dis. 2019; 21: e13005. [CrossRef]
  138. Cain O, Yoong A, Lipkin G, Huengsberg M, Murray J, Rudzki Z, et al. Rapidly progressive intravascular primary effusion lymphoma in an HIV-positive renal transplant recipient. Histopathology. 2018; 72: 339-341. [CrossRef]
  139. Patel A, Bishburg E, Zucker M, Tsang P, Nagarakanti S, Sabnani I. Concomitant Kaposi sarcoma and multicentric Castleman's disease in a heart transplant recipient. Heart Lung. 2014; 43: 506-509. [CrossRef]
  140. Mandel C, Silberstein M, Hennessy O. Case report: Fatal pulmonary Kaposi's sarcoma and Castleman's disease in a renal transplant recipient. Br J Radiol. 1993; 66: 264-265. [CrossRef]
  141. Gaitonde S, Vidanovic V, Ni H. Concomitant and fatal HHV-8+ multicentric Castleman's disease and Kaposi's sarcoma in the same lymph node of an HIV- liver transplant patient. Histopathology. 2007; 50: 954-958. [CrossRef]
  142. Vijgen S, Wyss C, Meylan P, Bisig B, Letovanec I, Manuel O, et al. Fatal outcome of multiple clinical presentations of human herpesvirus 8-related disease after solid organ transplantation. Transplantation. 2016; 100: 134-140. [CrossRef]
  143. Karras A, Thervet E, Legendre C, Groupe Coopératif de transplantation d'Ile de France. Hemophagocytic syndrome in renal transplant recipients: Report of 17 cases and review of literature. Transplantation. 2004; 77: 238-243. [CrossRef]
  144. Luppi M, Barozzi P, Rasini V, Riva G, Re A, Rossi G, et al. Severe pancytopenia and hemophagocytosis after HHV-8 primary infection in a renal transplant patient successfully treated with foscarnet. Transplantation. 2002; 74: 131-132. [CrossRef]
  145. Luppi M, Barozzi P, Schulz TF, Setti G, Staskus K, Trovato R, et al. Bone marrow failure associated with human herpesvirus 8 infection after transplantation. N Engl J Med. 2000; 343: 1378-1385. [CrossRef]
  146. Pietrosi G, Vizzini G, Pipitone L, Di Martino G, Minervini MI, Lo Iacono G, et al. Primary and reactivated HHV8 infection and disease after liver transplantation: A prospective study. Am J Transplant. 2011; 11: 2715-2723. [CrossRef]
  147. Cohen GM, Langer AL, Sima H, Chang C, Troy K, Taimur S. Hemophagocytic lymphohistiocytosis due to primary HHV-8 infection in a liver transplant recipient. Transplant Direct. 2018; 4: e411. [CrossRef]
  148. Gantt S, Casper C. Human herpesvirus 8-associated neoplasms: The roles of viral replication and antiviral treatment. Curr Opin Infect Dis. 2011; 24: 295-301. [CrossRef]
  149. Riva G, Luppi M, Barozzi P, Forghieri F, Potenza L. How I treat HHV8/KSHV-related diseases in posttransplant patients. Blood. 2012; 120: 4150-4159. [CrossRef]
  150. Chiereghin A, Barozzi P, Petrisli E, Piccirilli G, Gabrielli L, Riva G, et al. Multicenter prospective study for laboratory diagnosis of hhv8 infection in solid organ donors and transplant recipients and evaluation of the clinical impact after transplantation. Transplantation. 2017; 101: 1935-1944. [CrossRef]
  151. Gantt S, Cattamanchi A, Krantz E, Magaret A, Selke S, Kuntz SR, et al. Reduced human herpesvirus-8 oropharyngeal shedding associated with protease inhibitor-based antiretroviral therapy. J Clin Virol. 2014; 60: 127-132. [CrossRef]
  152. Spira TJ, Lam L, Dollard SC, Meng YX, Pau CP, Black JB, et al. Comparison of serologic assays and PCR for diagnosis of human herpesvirus 8 infection. J Clin Microbiol. 2000; 38: 2174-2180.
  153. Laney AS, Peters JS, Manzi SM, Kingsley LA, Chang Y, Moore PS. Use of a multiantigen detection algorithm for diagnosis of Kaposi's sarcoma-associated herpesvirus infection. J Clin Microbiol. 2006; 44: 3734-3741. [CrossRef]
  154. Engels EA, Biggar RJ, Marshall VA, Walters MAKL, Gamache CJ, Whitby D, et al. Detection and quantification of Kaposi's sarcoma-associated herpesvirus to predict AIDS-associated Kaposi's sarcoma. AIDS. 2003; 17: 1847-1851. [CrossRef]
  155. Whitby D, Howard MR, Tenant-Flowers M, Brink NS, Copas A, Boshoff C, et al. Detection of Kaposi sarcoma associated herpesvirus in peripheral blood of HIV-infected individuals and progression to Kaposi's sarcoma. Lancet. 1995; 346: 799-802. [CrossRef]
  156. Marcelin AG, Motol J, Guihot A, Caumes E, Viard JP, Dussaix E, et al. Relationship between the quantity of Kaposi sarcoma-associated herpesvirus (KSHV) in peripheral blood and effusion fluid samples and KSHV-associated disease. J Infect Dis. 2007; 196: 1163-1166. [CrossRef]
  157. Pellet C, Chevret S, Francès C, Euvrard S, Hurault M, Legendre C, et al. Prognostic value of quantitative Kaposi sarcoma--associated herpesvirus load in posttransplantation Kaposi sarcoma. J Infect Dis. 2002; 186: 110-113. [CrossRef]
  158. Ariza-Heredia EJ, Razonable RR. Human herpes virus 8 in solid organ transplantation. Transplantation. 2011; 92: 837-844. [CrossRef]
  159. Luppi M, Barozzi P, Guaraldi G, Ravazzini L, Rasini V, Spano C, et al. Human herpesvirus 8-associated diseases in solid-organ transplantation: Importance of viral transmission from the donor. Clin Infect Dis. 2003; 37: 606-607. [CrossRef]
  160. Serraino D, Piselli P, Scuderi M, Gabbrielli F, Venettoni S, Grossi P, et al. Screening for human herpesvirus 8 antibodies in Italian organ transplantation centers. Clin Infect Dis. 2005; 40: 203-205. [CrossRef]
  161. Barozzi P, Bonini C, Potenza L, Masetti M, Cappelli G, Gruarin P, et al. Changes in the immune responses against human herpesvirus-8 in the disease course of posttransplant Kaposi sarcoma. Transplantation. 2008; 86: 738-744. [CrossRef]
  162. Campistol JM, Schena FP. Kaposi's sarcoma in renal transplant recipients--the impact of proliferation signal inhibitors. Nephrol Dial Transplant. 2007; 22: i17-i22. [CrossRef]
  163. Detroyer D, Deraedt K, Schöffski P, Hauben E, Lagrou K, Naesens M, et al. Resolution of diffuse skin and systemic Kaposi's sarcoma in a renal transplant recipient after introduction of everolimus: A case report. Transpl Infect Dis. 2015; 17: 303-307. [CrossRef]
  164. Fu W, Merola J, Malinis M, Lacy J, Barbieri A, Liapakis AH, et al. Successful treatment of primary donor-derived human herpesvirus-8 infection and hepatic Kaposi sarcoma in an adult liver transplant recipient. Transpl Infect Dis. 2018; 20: e12966. [CrossRef]
  165. Stallone G, Schena A, Infante B, Di Paolo S, Loverre A, Maggio G, et al. Sirolimus for Kaposi's sarcoma in renal-transplant recipients. N Engl J Med. 2005; 352: 1317-1323. [CrossRef]
  166. Stallone G, Infante B, Grandaliano G, Schena FP, Gesualdo L. Kaposi's sarcoma and mTOR: A crossroad between viral infection neoangiogenesis and immunosuppression. Transplant Int. 2008; 21: 825-832. [CrossRef]
  167. Guba M, von Breitenbuch P, Steinbauer M, Koehl G, Flegel S, Hornung M, et al. Rapamycin inhibits primary and metastatic tumor growth by antiangiogenesis: Involvement of vascular endothelial growth factor. Nat Med. 2002; 8: 128-135. [CrossRef]
  168. Nichols LA, Adang LA, Kedes DH. Rapamycin blocks production of KSHV/HHV8: Insights into the anti-tumor activity of an immunosuppressant drug. PloS one. 2011; 6: e14535. [CrossRef]
  169. Neyts J, De Clercq E. Antiviral drug susceptibility of human herpesvirus 8. Antimicrob Agents Chemother. 1997; 41: 2754-2756. [CrossRef]
  170. Casper C, Krantz EM, Corey L, Kuntz SR, Wang J, Selke S, et al. Valganciclovir for suppression of human herpesvirus-8 replication: A randomized, double-blind, placebo-controlled, crossover trial. J Infect Dis. 2008; 198: 23-30. [CrossRef]
  171. Kowalkowski MA, Kramer JR, Richardson PR, Suteria I, Chiao EY. Use of boosted protease inhibitors reduces Kaposi sarcoma incidence among male veterans with HIV infection. Clin Infect Dis. 2015; 60: 1405-1414. [CrossRef]
  172. Chang E, Mapakshi SR, Mbang P, El-Mallawany NK, Kramer JR, White DL, et al. Impact of protease inhibitors on HIV-associated kaposi sarcoma incidence: A systematic review. J Acquir Immune Defic Syndr. 2018; 79: 141-148. [CrossRef]
  173. Sgadari C, Barillari G, Toschi E, Carlei D, Bacigalupo I, Baccarini S, et al. HIV protease inhibitors are potent anti-angiogenic molecules and promote regression of Kaposi sarcoma. Nat Med. 2002; 8: 225-232. [CrossRef]
  174. Gantt S, Carlsson J, Ikoma M, Gachelet E, Gray M, Geballe AP, et al. The HIV protease inhibitor nelfinavir inhibits Kaposi's sarcoma-associated herpesvirus replication in vitro. Antimicrob Agents Chemother. 2011; 55: 2696-2703. [CrossRef]
  175. Gantt S, Casper C, Ambinder RF. Insights into the broad cellular effects of nelfinavir and the HIV protease inhibitors supporting their role in cancer treatment and prevention. Curr Opin Oncol. 2013; 25: 495-502. [CrossRef]
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