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Clinical Transplantation

Infectious Enteritis After Intestinal Transplantation: Incidence, Timing, and Outcome

Ziring, David2; Tran, Robert2; Edelstein, Susan2; McDiarmid, Sue V.2; Gajjar, Nupoor3; Cortina, Galen3; Vargas, Jorge2; Renz, John F.1; Cherry, James D.2; Krogstad, Paul2; Miller, Marjorie3; Busuttil, Ronald W.1; Farmer, Douglas G.1,4

Author Information
doi: 10.1097/01.TP.0000154911.15693.80


Since the 1990s, the field of intestinal transplantation (ITx) has seen major advancements that have translated into improved clinical outcomes (1–7). Without a doubt, advances in immunosuppression have played a significant role (8). However, immune therapy has always been a double-edged sword in organ transplantation: Too little or inappropriate therapy may result in allograft rejection, whereas too much therapy may facilitate infections. ITx has been particularly prone to this principle because the intestine is susceptible to rejection and injury to the intestine can result in translocation of infectious organisms. This problem has been illustrated in the data from the International Intestinal Transplant Registry, which show that the leading cause of patient death after ITx is sepsis and that acute cellular rejection (ACR) rates are reported to be as high as 79% with resulting graft loss a common problem (1).

As the field of ITx has advanced, there have been reports documenting intestinal infections (7,9–13). Furthermore, because the intestine is a frequent target for cytomegalovirus (CMV) and Epstein-Barr virus (EBV) infection (14), it is not surprising that CMV and EBV have been problematic with disease rates as high as 40%. No doubt, other gastrointestinal infectious agents have affected ITx recipients, many of which have not been reported or diagnosed. A common clinical scenario is the ITx recipient with abnormal allograft function, fevers, and abnormal intestinal biopsy. The differential diagnosis includes ACR, but many infections, particularly viral, can mimic this clinicopathologic picture. Because the treatments for infectious enteritis (IE) and ACR are polar opposites, the timely, accurate distinction between these two entities is of critical importance. Furthermore, it is also possible that the presence of IE may predispose patients to ACR, thereby increasing the potential morbidity and mortality associated with this process (15).

Faced with this clinical dilemma, we retrospectively reviewed our experience with IE after ITx. This study identifies for the first time the wide range of infectious agents that has been encountered after ITx, explores the relationship between IE and rejection, and develops a diagnostic and therapeutic plan based on lessons learned from this experience.



Institutional Review Board approval was obtained for this study in compliance with Health Insurance Portability and Accountability Act guidelines. With the use of a transplant database and hospital medical records, we included all patients who underwent ITx between November 1991 and May 2003 at the Dumont-UCLA Transplant Center. Follow-up was concluded in August 2003. The details regarding demographics, types of transplants, postoperative immunosuppression, and outcomes were recorded. Particular focus was placed on posttransplant microbiology, and histopathology. IE was defined as the presence of clinical symptoms of abnormal allograft function associated with (1) abnormal histopathology and infectious agents identified, or (2) ostomy culture positive for an infectious agent.

Patient Management: General

All recipients of ITx allografts are cared for by a single transplant team under protocoled management as published elsewhere (6). Perioperative broad-spectrum antibacterial antibiotics are administered for 48 to 72 hr after ITx. Further antimicrobial therapy is dictated by the clinical course. Routine culture and sensitivity studies are not obtained except from the respiratory tract and blood pretransplant. Clinical signs of infection prompt an infectious workup that includes general bacterial, fungal, and viral culture of respiratory secretions, blood, urine, and stool/ostomy effluent. Specific details regarding gastrointestinal cultures are outlined next. Antimicrobial sensitivity patterns are determined on all relevant bacterial isolates and many fungal isolates.

Immunosuppression Regimens

As in most ITx centers, immunotherapeutic regimens have evolved throughout the experience. In short, all patients have been managed with tacrolimus-based (Prograf, Fujisawa, Deerfield, IL) triple immunosuppression. The first four patients also received induction orthoclone muromonab-CD3 (OKT3, Ortho Biotech Products, Raritan, NJ). Our most recent standard immunosuppression regimen that has been systematically applied since 1999 includes induction interleukin-2 receptor antagonist with daclizumab (Zenapax, Roche Laboratories, Nutley, NJ) or basiliximab (Simulect, Novartis Pharmaceuticals, East Hanover, NJ) combined with tacrolimus, corticosteroids, and mycophenolate mofetil (CellCept, Roche Laboratories). Target tacrolimus trough levels were between 10 and 20 ng/mL. Sirolimus (Rapamune, Wyeth-Ayerst Pharmaceuticals, Inc., Philadelphia, PA) was used as a rescue agent when the tacrolimus-based immunotherapies failed. ACR was diagnosed by histopathology and treated with high-dose steroids or OKT3.

Antiviral Prophylaxis Regimens

All recipients and most donors are tested pretransplant for the presence of antibodies against CMV and EBV. No effort has been made to match CMV or EBV donor:recipient combinations. A standard antiviral prophylactic regimen has been used since 1996 based on our experience in liver transplant recipients (16) and published elsewhere (17). Briefly, the regimen consists of intravenous ganciclovir (Cytovene, Roche Laboratories) administered at a dose of 10 mg/kg per day for the first 14 days and 6 mg/kg per day for the next 86 days. Thereafter, patients are converted to and maintained on prophylactic oral acyclovir (Zovirax, GlaxoSmithKline, Research Triangle Park, NC) at 40 mg/kg per day in divided doses for an indefinite span of time. Consideration is given toward discontinuing the acyclovir after 5 years of a successful ITx.

In addition, preemptive therapy has been used in all patients since 1996. Peripheral blood is routinely tested for the presence of CMV (qualitative polymerase chain reaction [PCR] or quantitative hybridization antibody capture assay [Digene, Inc., Gaithersburg, MD]) or EBV DNA by quantitative PCR. Positive assays prompt preemptive therapy with ganciclovir alone or in combination with CMV immune globulin (Cytogam, MedImmune, Gaithersburg, MD). Recipients with high EBV DNA copy numbers or suspicious clinical symptoms were evaluated for the presence of posttransplant lymphoproliferative disorders (PTLDs) by radiographic scans and tissue biopsies.

Intestinal Surveillance

Protocol biopsies of the transplanted intestine are generally obtained once weekly starting on postoperative days 10 to 14 and concluding 8 weeks after ITx. Endoscopy and biopsies are also obtained when clinical symptoms indicating abnormal allograft function are present. Multiple targeted biopsies are obtained and sent to microbiology for general viral culture and to pathology for routine hematoxylin-eosin (H&E) staining. In addition, since 1999, histopathology samples also undergo immunostaining with probes targeted against CMV, herpes simplex viruses I and II, and adenovirus. Abnormal results prompt further endoscopic surveillance as indicated.

Infectious Disease Evaluation

Routine screening cultures have not been used. Evaluation is prompted by changes in clinical symptoms and allograft function. Endoscopy and biopsy are undertaken, and ostomy effluent or stool is sent for a clinically relevant battery of tests. The extent of this testing has evolved over time and is tailored to the individual patient situation. The battery can include the following: general bacterial culture; general fungal culture; general viral culture; shell vial assay, cell count, and differential cell count; ova and parasite examination; Giardia lamblia direct immunofluorescence assay (Merifluor Detection exam, Meridian Bioscience, Inc., Cincinnati, OH); Cryptosporidium direct immunofluorescence assay (Merifluor Detection exam, Meridian Bioscience, Inc.); Isospora modified Ziehl-Neelsen acid-fast stain (Medical Chemical Corp., Torrance, CA); Clostridium difficile toxin A and B enzyme immunoassay (Meridian Bioscience, Inc.); and rotavirus antigen enzyme immunoassay (Premier Rotaclone, Meridian Bioscience, Inc.). For general viral culture, stool samples are inoculated into three cell lines for tube culture and two shell vials (Diagnostic Hybrids, Athens, OH). The tube cultures are maintained for 3 weeks and observed for the development of cytopathic effect. Shell vials are incubated for 48 hr; one vial is stained with direct fluorescent antibody with a pool containing seven monoclonal antibodies (to adenovirus group antigen, respiratory syncytial virus, influenza A and B, and parainfluenza types 1, 2, and 3 [Diagnostic Hybrids]). If positive, the second vial is stained with the individual antisera for final identification. CMV is detected similarly by inoculating cell lines in both tube culture and shell vial, and positive cultures are detected using direct fluorescent antibody (Chemicon, Temecula, CA) to CMV antigen. Tube cultures are maintained for 4 weeks, whereas CMV shell vial culture is stained at 48 hr. Cultures and antigen detection methods for adenovirus 40 and 41 were not routinely used. Clinical suspicion may also result in blood cultures, urine cultures, and CMV and EBV DNA testing as described.


Survival was calculated with the Kaplan-Meier method. The log-rank test was used to compare survival differences. The median follow-up of the study group was 12 (2–69) months.



During the study interval, 33 patients underwent 37 ITxs. Thirteen of these recipients (39%) developed IE and are included in the analysis. There were 20 episodes of IE in these 13 recipients. Eight (IE+) patients had one episode, four patients had two episodes, and one patient had four episodes. The demographics of the study population with IE included three adults (>18 years) who were male. There were 10 children (≤18 years) (77% of study group), including seven males, and three females. The study population had a median age of 34 (10–585) months. Among the three adults who developed IE, the causes of their infections were CMV, G. lamblia, and rotavirus. Of the 20 other patients who underwent ITx but did not develop IE during the study period (IE−), nine were adults (four males and five females) and 11 were children (four males and seven females) (55%).

All recipients in the study group with IE initially had intestinal failure and were dependent on total parenteral nutrition. Their diagnoses were necrotizing enterocolitis (most frequent), gastroschisis, midgut volvulus, microvillus inclusion disease, Hirschsprung disease, inflammatory bowel disease, jejunoileal atresia, and pseudo-obstruction. The majority of these diagnoses were nonmotility disorders, and only one patient required ITx secondary to pseudo-obstruction. Eleven patients with IE received combined liver plus intestinal grafts, whereas two received isolated intestinal grafts. Immunotherapy was tacrolimus-based in all patients with quadruple therapy with interleukin-2 receptor antagonist (8). The overall 1- and 3-year actuarial patient and graft survivals are shown in Figure 1A and B. Compared with the cohort of recipients without infections, recipients with IE demonstrated a slight but not significant survival disadvantage. The two intestinal allograft losses were associated directly with viral IE. Of the 33 total patients who underwent ITx, four patients (all males, two adults) required regrafting. Two of these patients (one adult, one child) developed IE.

(A) Patient survival: 36-month patient survival was 57% for IE+ and 52% for IE− patients. Differences were not statistically significant. (B) Graft survival: 36-month graft survival was 53% for IE+ and 37% for IE− patients. Differences were not statistically significant.

The median time from ITx to the first episode of IE was 76 (32–1,800) days. Seventy percent of the IE episodes were viral in cause, whereas 15% each were bacterial or parasitic (Fig. 2). Fourteen (70%) episodes of IE were diagnosed in patients hospitalized within the incubation period of the infection and therefore were likely nosocomially acquired.

Infectious agents causing IE: 70% were viral, 15% were bacterial, and 15% were parasitic. The most common overall pathogen was rotavirus. EBV, Epstein-Barr virus; CMV, cytomegalovirus.

Thirty percent of the episodes of IE were associated with ACR. The median time from diagnosis of IE to ACR was 36 (0–52) days. The agents most often associated with ACR were adenovirus, rotavirus, and Cryptosporidium.

Viral Enteritis

Twelve patients experienced 14 episodes of viral IE. The viral agents were rotavirus (six patients/eight infections; five pediatric; five male), adenovirus (four patients/four infections; four pediatric; two male) (Fig. 3A and B), CMV (one adult male patient/one infection) (Fig. 4), and EBV (one pediatric female patient/one infection). The rotavirus infections were all diagnosed with stool rotavirus antigen enzyme assay and occurred a median of 75 (33–1,800) days after ITx. Supportive care, including intravenous fluid and parenteral nutrition, was the only treatment instituted. One graft was lost to ACR that followed a rotaviral infection. Prolonged viral carriage as long as 6 weeks as documented by persistent symptoms and positive rotaviral antigen testing occurred frequently.

Viral IE after ITx. (A) Adenovirus. Hematoxylin-eosin (H&E)-stained section (400× magnification) with adenoviral inclusion (arrow). (B) Immunoperoxidase-stained section highlighting adenovirus positive staining cells (arrow) (400× magnification).
CMV enteritis after ITx on H&E-stained tissue demonstrating CMV viral inclusions (arrow).

Adenoviral IE occurred a median of 113 (32–541) days after ITx. All episodes were accompanied by abnormal histopathology that resembled ACR with increased apoptosis, lymphocytic infiltration, and architectural damage. Three of these infections were initially misdiagnosed and treated with steroid bolus for ACR. In two patients, the correct diagnosis was obtained through immunostaining between 3 and 5 days after initiation of ACR treatment (Fig. 3B). ACR therapy was immediately stopped, and antiviral therapy with ribavirin (ICN Pharmaceuticals Inc., Costa Mesa, CA) and intravenous immunoglobulin was initiated. Complete resolution was seen in both patients. The third case was not correctly diagnosed until after the allograft was removed for what was thought to be refractory ACR. Then, the initial viral stool cultures obtained at the onset of symptoms became positive, and retrospective staining of the initial biopsies and explant were also positive for adenovirus. The last case occurred in a symptomatic recipient with allograft dysfunction from an unknown cause. No treatment was administered, and the clinical symptoms resolved. Stool viral cultures returned positive for adenovirus more than 30 days after they were obtained.

Only two cases of tissue-invasive CMV or EBV disease were noted. EBV disease was diagnosed by Epstein-Barr virus encoded small RNA stain on a biopsy in a child 476 days after ITx who was undergoing OKT3 therapy for steroid-resistant ACR. Rejection therapy was completed, ganciclovir and CMV immunoglobulin were administered, and the allograft dysfunction resolved. The lone case of CMV disease in an intestinal allograft was diagnosed on histopathology 1,467 days after ITx (Fig. 4). The infection was treated with intravenous ganciclovir and CMV immunoglobulin.

Parasitic Enteritis

Three patients developed parasitic IE that occurred a median of 183 (51–298) days after ITx. The infectious agents were G. lamblia (one adult male patient/one infection) (Fig. 5) and Cryptosporidium sp. (two patients/two infections; two pediatric; one male) (Fig. 6). Giardiasis and one episode of cryptosporidiosis were nosocomially acquired. Giardiasis occurred 183 days after ITx, was diagnosed on H&E histopathology staining, and resolved with metronidazole (Flagyl, Searle, Skoke, IL) therapy. Both Cryptosporidium infections were diagnosed by stool culture or histopathology. Both cases were successfully treated with prolonged courses of paromomycin (Humatin, Parke Davis, Ann Arbor, MI) and azithromycin (Zithromax, Pfizer Labs, New York, NY). ACR occurred almost simultaneously with the diagnosis of cryptosporidiosis in one case and was successfully treated with steroids simultaneously.

Parasitic IE after ITx on H&E-stained tissue. Giardia lamblia organisms (arrows).
Cryptosporidium trophozoites (arrows) in an ITx recipient.

Bacterial Enteritis

C. difficile was the only bacterial-related IE seen (two patients/three episodes; two pediatric; one male). All episodes were accompanied by clinical symptoms and, in cases in which biopsies were performed, there were minimal histopathologic changes. All were successfully treated with intravenous metronidazole or vancomycin (Vancocin, Eli Lilly and Co., Indianapolis, IN).


The success of ITx over the last decade has, in large part, been the result of advances in immunotherapy. However, infections are a common complication seen in immunosuppressed patients. Our experience with this complex group of recipients has identified a significant problem related to ITx: IE. Until now, only sporadic reports in the literature describing this complication exist (10–13,18–20). On the basis of our own experience, we believe that IE after ITx has been underdiagnosed, potentially incorrectly treated, and underreported. This study spanning more than a decade of experience represents one of the largest investigations to date on this subject and reveals several important issues.

First, our awareness of and ability to diagnose IE have markedly improved. This is borne out by the fact that IE was almost never diagnosed in the first 10 recipients but has been diagnosed with increasing frequency in the later experience. This increasing frequency of diagnosis is mainly because of clinical awareness. As our experience has grown, we have included a more systematic approach to diagnosing IE, which includes a routine battery of infectious diagnostic testing performed when patients become symptomatic. Therefore, the quoted 39% incidence of IE may in fact be misleading and could be higher. The true incidence will be revealed as more experience is gained with the newer, more extensive diagnostic protocols.

A second interesting finding is the preponderance of IE in male children undergoing ITx. Although our overall numbers are small, thereby limiting statistical interpretation, we cannot ignore the fact that 77% of those with IE were male and 85% were children. For comparison, in the 20 other recipients who underwent transplantation during the study interval who were not diagnosed with IE, 40% were male and 55% were children. This gender predilection of IE is not surprising, because the incidence and severity of many infectious diseases, including gastroenteritis, is known to be higher in males.

In support of this concept is the fact that IE is not nearly as common after other comparable transplants such as the liver (21,22). Therefore, several unique aspects of the intestinal allograft are worthy of mention. First is the fact that many ITx donors are children (79% of ITx donors in 2003) (23), and the intestines of children may have a more naïve or immature GALT. Second is the fact that the immune effector cells within the intestinal allograft are rapidly replaced by recipient-derived effector cells (24) shortly after transplantation, exposing the possibility of an acquired GALT immunodeficiency. It is possible that the mixture of the human leukocyte antigen (HLA)-mismatched recipient immune effector cells and donor stromal antigen-presenting cells creates a less-efficient local environment for an immune response. To our knowledge, however, there are no data to support this argument. The only data regarding the immunocompetency of intestinal allografts include a single study examining surface immunoglobulin-A responses to cholera toxin after rat ITx (25), showing an impaired surface immunoglobulin-A response. Clearly, further investigation is warranted.

Complicating this immune environment is the relative impact of various immunotherapeutic regimens. Without a doubt, ITx recipients require significant levels of immunosuppression. Whether the specific regimens used in this patient population directly contribute to the incidence of IE is unknown and cannot be determined by a study of this nature. With our quadruple induction immunotherapy protocol, we seek to reduce the overall immunosuppression by reducing doses of each agent. It is not possible to quantify the overall state of immunosuppression in these patients, and therefore we cannot determine this important variable. Again, this is another area worthy of investigation.

The fact that IE can be associated with ACR should not be surprising. As noted, the cells responsible for infectious surveillance are likely recipient derived and therefore of a different HLA type than the surrounding stromal and antigen-presenting cells. Because presentation of infectious antigens such as viral antigens occurs in the context of HLA class I molecules, it is a matter of semantics whether the recipient T cells are responding to the viral epitopes presented by foreign HLA antigens or are responding to foreign HLA antigen epitopes themselves (26). In either context, the immune response is similar, thus explaining the difficulty in distinguishing IE, particularly viral in origin, from ACR. Our experience supports these theoretic concerns. That is, with the exception of one episode, all ACR episodes followed viral infections.

This experience has exposed the need for a rapid and accurate method of diagnosing the presence of viral enteritis. We believe this is of critical importance because successful treatment of IE or ACR requires the prompt institution of opposite therapies. We address this issue systematically. First, when allograft dysfunction occurs, a stool sample is sent to microbiology for general viral culture. Our laboratory uses a shell vial assay with probes directed against specific viral antigens that results in detection of adenovirus, CMV, and EBV, usually within 48 hr (27). An excellent alternative to this assay, providing rapid, accurate results, is PCR to detect specific viral genome sequences, as championed by others (18). Second, our pathology laboratory routinely processes biopsy tissue by performing immunostains against these viruses and provides results within 48 hr. Third, we do not treat allograft dysfunction before obtaining tissue biopsy and undertaking the special infectious investigation as outlined except under unusual circumstances. Combining prompt infectious workup with biopsy will afford ample time to institute the proper therapy to correctly treat and rescue the transplanted allograft. This is of critical importance because in this experience, one allograft was lost to adenovirus enteritis that was incorrectly treated as ACR.

The majority of our patients with IE were children. Children undergoing ITx may be at increased risk of developing common childhood viral illnesses (e.g., rotavirus) because of the generally high incidence of these infections in the general pediatric population. The most common viral infections seen in this patient population were adenovirus and rotavirus. Although adenoviral enteritis after ITx has been reported before, this represents the first report of rotaviral enteritis after ITx. In general, rotavirus can be accurately diagnosed in minutes with the use of rapid enzyme-linked immunosorbent assay (28). It was most frequently diagnosed in the fall, winter, and spring (four of eight cases), consistent with its natural cyclical incidence. Although rotavirus enteritis is generally assumed to be a pediatric disease, one of the six patients with this infection was an adult. Our treatment for patients with rotaviral infection is to provide supportive care when necessary and to survey the intestine by endoscopy and biopsy while the patient is recovering. Most of these infections are self-limited, but it is possible to develop ACR during or after rotaviral infection, and this should be promptly diagnosed and treated.

Adenovirus infection is more difficult and concerning. Adenovirus infections as a complication of liver transplantation, particularly among children, have been reported with mortality rates as high as 10% (21,22). Its incidence after ITx is probably incompletely reported. The existing literature includes case reports (10,19,20) and a recent series with an incidence of 26% (12). When the intestine is involved, the response to adenovirus infection typically includes histopathologic changes of crypt cell apoptosis and nuclear disarray, eosinophilic nuclear inclusions, “smudge cells” with enlarged basophilic nuclei, surface enterocyte proliferation, and an inflammatory infiltrate that is primarily plasmacytic in origin (29). These changes cannot be relied on for diagnosis because many overlap with those seen in rejection (18,29). Because some adenoviruses are difficult to grow in culture, we strongly recommend the use of rapid diagnostic tests such as shell vial culture with immunostains, as well as enzyme immunoassay specifically for serotypes 40 and 41, for diagnosis. Other rapid and sensitive methods to detect adenoviruses such as PCR and electron microscopy do exist and can be considered (10,12,18). Treatment should include supportive care and reduction in immunosuppression. There are no uniformly effective, accepted treatments for adenoviruses. Intravenous ribavirin has been used with limited success in some bone marrow transplant recipients (30), and less frequently in solid-organ transplant recipients (12,31,32). Newer antivirals such as cidofovir have shown success in case studies (18,33–35). We believe that the use of both ribavirin and intravenous immunoglobulin may be helpful in the treatment of these patients. As shown by our experience, successful treatment with complete resolution and full graft function is possible in the majority of cases, although increased morbidity and mortality have been reported by others (12,19).

Typically, CMV and EBV disease including PTLD have been a common occurrence after ITx (1,2,4). In contrast, the incidence of CMV and EBV infection is low in our patient population as is the incidence of graft loss from these viruses or the development of PTLD. We attribute this to an aggressive protocol that involves both prophylactic and preemptive therapies directed against these viruses (17). Others have recommended similar treatment and prophylactic strategies (36). The success of these protocols has allowed the use of CMV-positive donor organs in all recipients.

Other viral infections are certainly possible, although they were not diagnosed in this population. A good argument can be made to institute rapid assay diagnostic tests for enteroviruses and calciviruses, because they have been diagnosed after ITx by other groups (13).

Cryptosporidium enteritis has been reported in the literature in five ITx recipients (9,11). In our experience, cryptosporidial organisms were easily diagnosed on H&E staining of intestinal biopsies and by direct immunofluorescent stain of stool samples. There are now several treatment options available for Cryptosporidium. We demonstrated successful and complete resolution of these infections with the combination use of paromomycin and azithromycin as is recommended in other immunosuppressed conditions (37). The average length of antibiotic treatment required to clear the infection was 58 days. One infection was associated with ACR, but others have not noted this association (9,11). Although we had success with treatment of both of our patients, immunocompromised patients with cryptosporidiosis historically show an inconsistent response to paromomycin and azithromycin combination therapy (38). Newer agents have been introduced, but the few studies using nitazoxanide (Alinia, Romark Laboratories, Tampa, FL) in immunosuppressed children (those with acquired immune deficiency syndrome) show no advantage (39). In an effort to prevent the development of this infection, we recommend that the use of untreated tap and well water be avoided for all enteral intake (40).

This is the first report of intestinal giardiasis after ITx. This infection was easily diagnosed on histopathology and was also detected using microbiologic assay. Treatment was successful with metronidazole, and we follow the same recommendations as for Cryptosporidium to prevent infections.

Finally, the major bacterial-related infection encountered was C. difficile. This again represents the first report of this infection after ITx. It is not surprising that this type of infection, which develops usually as a consequence of antimicrobial therapy, was seen in this patient population. All cases were accurately diagnosed using microbiologic enzyme immunoassay for toxins A or B. Standard supportive care as outlined is necessary, and the standard therapeutic options seem to be applicable to this patient population.


We present one of the first reports on a broad range of IE after ITx. There was a preponderance of these types of infections in male children after ITx for unexplained reasons. Viral IE was the most serious infection, and approximately one third of infections were associated with rejection. Most infections responded to appropriate therapy, although two allografts were lost as a direct result of infection. On the basis of our experience, we offer the following recommendations:

  1. Institute surveillance including endoscopy, biopsy, and microbiologic assays promptly when allograft dysfunction occurs.
  2. Establish a policy for expeditious handling and microbiologic processing of all samples.
  3. Perform rapid diagnostic testing such as PCR, enzyme-linked immunosorbent assay, immunohistochemical staining, or shell vial assays for the diagnosis of common infections such as rotaviruses and adenoviruses.
  4. Send a portion of all biopsy samples to microbiology for culture and assay.
  5. Avoid the initiation of rejection therapy until infectious causes have been ruled out.
  6. During therapy for rejection or infection, assess the response to therapy frequently and periodically perform the same diagnostic assays not only to confirm therapeutic efficacy but also to ensure that an alternate process has not developed in the interim.


1. Grant D. Intestinal transplantation: 1997 Report of The International Registry. Transplantation 1999; 67: 1061–1064.
2. Abu-Elmagd K, Reyes J, Bond G, et al. Clinical intestinal transplantation: a decade of experience at a single center. Ann Surg 2001; 234: 404–416.
3. Nishida S, Levi D, Kato T, et al. Ninety-five cases of intestinal transplantation at the University of Miami. J Gastrointest Surg 2002; 6: 233–239.
4. Langnas A, Chinnakotla S, Sudan D, et al. Intestinal transplantation at the University of Nebraska Medical Center: 1990 to 2001. Transplant Proc 2002; 34: 958–960.
5. Farmer DG, McDiarmid SV, Yersiz H, et al. Outcomes after intestinal transplantation: a single-center experience over a decade. Transplant Proc 2002; 34: 896–897.
6. Farmer DG, McDiarmid SV, Yersiz H, et al. Outcome after intestinal transplantation: results from one center's 9-year experience; discussion 1031–2. Arch Surg 2001; 136: 1027–1031.
7. Fishbein TM, Florman S, Gondolesi G, et al. Intestinal transplantation before and after the introduction of sirolimus. Transplantation 2002; 73: 1538–1542.
8. Farmer DG. Clinical immunosuppression for intestinal transplantation. Curr Opin Organ Transplant 2004; 9: 214–219.
9. Gerber DA, Green M, Jaffe R, et al. Cryptosporidial infections after solid organ transplantation in children. Pediatr Transplant 2000; 4: 50–55.
10. Pinchoff RJ, Kaufman SS, Magid MS, et al. Adenovirus infection in pediatric small bowel transplantation recipients. Transplantation 2003; 76(1): 183–189.
11. Delis SG, Tector J, Kato T, et al. Diagnosis and treatment of Cryptosporidium infection in intestinal transplant recipients. Transplant Proc 2002; 34: 951–952.
12. McLaughlin GE, Delis S, Kashimawo L, et al. Adenovirus infection in pediatric liver and intestinal transplant recipients: utility of DNA detection by PCR. Am J Transplant 2003; 3: 224–228.
13. Kaufman SS, Chatterjee NK, Fuschino ME, et al. Calicivirus enteritis in an intestinal transplant recipient. Am J Transplant 2003; 3: 764–768.
14. Fishman JA, Rubin RH. Infection in organ-transplant recipients. N Engl J Med 1998; 338: 1741–1751.
15. Cainelli F, Vento S. Infections and solid organ transplant rejection: a cause-and-effect relationship? Lancet Infect Dis 2002; 2: 539–549.
16. Winston DJ, Wirin D, Shaked A, et al. Randomized comparison of ganciclovir and high-dose acyclovir for long-term cytomegalovirus prophylaxis in liver-transplant recipients. Lancet 1995; 346: 69–74.
17. Farmer DG, McDiarmid SV, Winston D, et al. Effectiveness of aggressive prophylactic and preemptive therapies targeted against cytomegaloviral and Epstein-Barr viral disease after human intestinal transplantation. Transplant Proc 2002; 34: 948–949.
18. Kaufman SS, Magid MS, Tschernia A, et al. Discrimination between acute rejection and adenoviral enteritis in intestinal transplant recipients. Transplant Proc 2002; 34: 943–945.
19. Berho M, Torroella M, Viciana A, et al. Adenovirus enterocolitis in human small bowel transplants. Pediatr Transplant 1998; 2: 277–282.
20. Levy MF, Crippin JS, Abbasoglu O, et al. Adenovirus infection of the human intestinal allograft: a case report. Transplant Proc 1996; 28: 2786–2787.
21. McGrath D, Falagas ME, Freeman R, et al. Adenovirus infection in adult orthotopic liver transplant recipients: Incidence and clinical significance. J Infect Dis 1998; 177: 459–462.
22. Michaels MG, Green M, Wald ER, et al. Adenovirus infection in pediatric liver-transplant recipients. J Infect Dis 1992; 165: 170–174.
23. Organ Procurement and Transplantation Network data of the U.S. Scientific Registry of Transplant Recipients and the Organ Procurement and Transplantation Network: Transplant Data 1988-March 2004. (2004, June 29). 2004. Rockville, MD, and Richmond, VA, HHS/HRSA/OSP/DOT and United Network for Organ Sharing.
24. Starzl TE, Demetris AJ. Transplantation tolerance, microchimerism, and the two-way paradigm. Theor Med Bioeth 1998; 19: 441–455.
25. Xia WY, Kirkman RL. Immune function in transplanted small intestine. Total secretory IgA production and response against cholera toxin. Transplantation 1990; 49: 277–280.
26. Cainelli F, Vento S. Infections and solid organ transplant rejection: a cause-and-effect relationship? Lancet Infect Dis 2002; 2(9): 539.
27. Athmanathan S, Bandlapally S, Rao GN. Comparison of the sensitivity of a 24 h-shell vial assay, and conventional tube culture, in the isolation of Herpes simplex virus-1 from corneal scrapings. BMC Clin Pathol 2002; 2: 1.
28. Rabenau H, Knoll B, Allwinn R, et al. Improvement of the specificity of enzyme immunoassays for the detection of rotavirus and adenovirus in fecal specimens. Intervirology 1998; 41: 55–62.
29. Parizhskaya M, Walpusk J, Mazariegos G, et al. Enteric adenovirus infection in pediatric small bowel transplant recipients. Pediatr Dev Pathol 2001; 4: 122–128.
30. Kapelushnik J, Or R, Delukina M, et al. Intravenous ribavirin therapy for adenovirus gastroenteritis after bone-marrow transplantation. J Pediatr Gastroenterol Nutr 1995; 21: 110–112.
31. Arav-Boger R, Echavarria M, Forman M, et al. Clearance of adenoviral hepatitis with ribavirin therapy in a pediatric liver transplant recipient. Pediatr Infect Dis J 2000; 19: 1097–1100.
32. Shetty AK, Gans HA, So S, et al. Intravenous ribavirin therapy for adenovirus pneumonia. Pediatr Pulmonol 2000; 29: 69–73.
33. Carter BA, Karpen SJ, Quiros RE, et al. Intravenous cidofovir therapy for disseminated adenovirus in a pediatric liver transplant recipient. Hepatology 2002; 36: 662A.
34. Ribaud P, Scieux C, Freymuth F, et al. Successful treatment of adenovirus disease with intravenous cidofovir in an unrelated stem-cell transplant recipient. Clin Infect Dis 1999; 28: 690–691.
35. Legrand F, Berrebi D, Houhou N, et al. Early diagnosis of adenovirus infection and treatment with cidofovir after bone marrow transplantation in children. Bone Marrow Transplant 2001; 27: 621–626.
36. Green M, Bueno J, Rowe D, et al. Predictive negative value of persistent low Epstein-Barr virus viral load after intestinal transplantation in children. Transplantation 2000; 70: 593–596.
37. Smith NH, Cron S, Valdez LM, et al. Combination drug therapy for cryptosporidiosis in AIDS. J Infect Dis 1998; 178: 900–903.
38. Hicks P, Zwiener RJ, Squires J, et al. Azithromycin therapy for Cryptosporidium parvum infection in four children infected with human immunodeficiency virus. J Pediatr 1996; 129: 297–300.
39. Amadi B, Mwiya M, Musuku J, et al. Effect of nitazoxanide on morbidity and mortality in Zambian children with cryptosporidiosis: a randomised controlled trial. Lancet 2002; 360: 1375–1380.
40. Chen XM, Keithly JS, Paya CV, et al. Current concepts: cryptosporidiosis. N Engl J Med 2002; 346: 1723–1731.

Intestinal transplantation; Infectious enteritis; Adenovirus; Cryptosporidium; Cytomegalovirus; Epstein-Barr virus

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