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Cytomegalovirus is a significant opportunistic pathogen in immuno-compromised patients. Primary infection, reactivation of latent virus, and reinfection are possible and are often clinically silent. The onset of infection is marked by spiking pyrexia, which may resolve in a few days. Its severity is parallel with the level of immunosuppression, and is greatest in bone marrow transplant (BMT) recipients and AIDS patients with low CD4+ T-cell counts. Solid organ transplant recipients, patients receiving immunosuppressive chemotherapy, and subjects with congenital immunodeficiencies may also be symptomatic (Landolfo et al., 2003).
Cytomegalovirus is one of the most important pathogens that infect (SOT) recipients and is associated with increased morbidity and mortality (Beam and Razonable, 2012). Solid organ transplant recipients are particularly susceptible to CMV-related disease due to the immunosuppression necessary to prevent organ rejection. Patients receiving T-cell depleting therapies are at the highest risk The major risk factor for CMV pneumonia is a CMV-seronegative transplant recipient receiving a CMV-seropositive organ. Because of the multiple human strains of CMV, seropositive organ recipients are at risk of re-infection with a different strain of virus. The clinical syndrome is then usually less severe than in primary infection and the onset of disease is often delayed to approximately 6-8 weeks post-transplantation (Alexopouloset al., 2012).
Clinical signs of HCMV infection in transplant recipients may be absent or severe, although severe infection is now less frequent as a result of better prophylaxis. HCMV is initially localized in the transplanted organ, but then spreads throughout the gastrointestinal tract and to the retina, skin, endometrium, lungs, and CNS. HCMV disease is more difficult to treat in BMT compared with SOT recipients, and HCMV pneumonia has a high mortality rate, despite the recent introduction of specific antiviral drugs (Landolfo et al., 2003). Lastly, an immunosuppressive syndrome often related to HCMV infection in the late post-transplant period is characterized by superinfection with bacteria, fungi, and protozoa, perhaps due to disturbance of both the humoral and cellular immune response by HCMV (Landolfo et al., 2003).
CMV can cause very serious infection in HIV infection (Barrett et al., 2012).
In the upper gastrointestinal tract, CMV has been isolated from oesophageal, gastric and duodenal ulcers. Patients with oesophageal disease may present with painful dysphagia. In the lower gastrointestinal tract, patients with CMV may present with diarrhoea due to colitis (Barrett et al., 2012).
CMV may cause disease in the peripheral and central nervous system (Barrett et al., 2012). Replication in the CNS of AIDS patients produces some of the symptoms observed in congenital infection, and is often followed by encephalopathy (Landolfo et al., 2003).
It is difficult to diagnose CMV infection in immunocompromised patients as it requires not only detection of virus but also determining whether CMV is causing disease. CMV shedding and viremia are common in patients with impaired cellular immunity even when disease due to CMV is not present (Jahan, 2010).
Rapid and sensitive technique for diagnosis of CMV infection is of vital importance for the management of immunocompromised patients. A number of rapid and sensitive methods have been developed. These includes DNA probe techniques (Spector and Vacqier, 1983) , Polymerase chain reaction (PCR) (Zipeto et al., 1992), CMV antigen detection in biopsies and bronchoalveolar lavage (Emanuel et al., 1986) and immunofluorescence technique for detection of CMV early antigens in cell (Vander et al., 1988). Moreover, an assay has been developed for CMV antigenemia based on the detection of CMV immediate early antigen (pp65 ) in circulating leucocytes (Boeckh et al., 1996).
Histopathology remains the reference standard for diagnosis of tissue-invasive CMV disease (Razonable and Hayden, 2013). CMV infection is indicated by cellular and nuclear enlargement (cytomegalic cells) and the presence of amphophilic to basophilic cytoplasmic inclusions (aggregates of CMV nucleoproteins that are produced during viral replication) (Eid et al., 2010). The severity of CMV infection can be assessed based on the degree of histological involvement (Mattes et al., 2000).
While these histopathologic findings are highly characteristic of CMV infection (Mattes et al., 2000), atypical features may be present and may overlap in appearance both with reactive changes and with inclusions of other intracellular viruses. Hence, the diagnosis can be confirmed further by in situ hybridization (ISH) or immunohistochemical (IHC) testing (Razonable and Hayden, 2013). The ISH uses CMV-specific cDNA probes that bind to viral DNA in the cellular materialÂ (Razonable and Hayden, 2013). Likewise, IHC uses monoclonal or polyclonal antibody against early CMV antigen (ChemalyÂ et al., 2004
Histopathology requires an invasive procedure to obtain tissue samples for testing (Eid et al., 2010). As a result, clinicians are often hesitant to perform it. Moreover, repeated biopsies cannot be performed serially to assess the response to treatment (Eid et al., 2010). Accordingly, many clinicians rely on the demonstration of CMV in the peripheral blood by Nucleic acid amplification testing (NAT) or antigen testing to support the clinical diagnosis of tissue-invasive CMV disease in patients with compatible clinical signs and symptoms (Razonable and Humar, 2013).
Serology relies on the sensitive detection of antibodies against CMV in the blood (Vauloup-Fellous et al., 2013). CMV immunoglobulin M (CMV-IgM) is initially secreted during early CMV infection, and the detection of CMV-IgM by serologic assays is indicative of active, acute, or recent infection. Weeks into the course of primary infection, CMV-IgG antibody is secreted, and this antibody persists for life. The detection of CMV-IgG is indicative of previous or past infection (Vauloup-Fellous et al., 2013).
Many different assays have been described and evaluated for the detection of CMV IgG antibodies. Among these are complement fixation, ELISA, anticomplement immunofluorescence, radioimmunoassay, and indirect hemagglutination (Ross et al., 2011).
Many different assays are available for IgM detection, but enzyme-linked immunosorbent assays (ELISAs) are the most widely used. Recombinant IgM assays using recombinant HCMV proteins and peptides have been developed in an attempt to standardize serological assays. However, studies have shown poor correlation of results obtained with different commercial kits for IgM testing. In addition, assays for IgM antibody lack specificity for primary infection because of false-positive results, because IgM can persist for months after primary infection, and because IgM can be positive in reactivated CMV infections (Ross et al., 2011).
Because of the limitations of the IgM assays, IgG avidity assays are utilized in some populations to help distinguish primary from non-primary CMV infection. These assays are based on the observation that IgG antibodies of low avidity are present during the first few months after the onset of infection and avidity increases over time reflecting maturation of the immune response. Thus, high anti-CMV IgG avidity represents longstanding infection in an individual. Avidity levels are reported as the avidity index which is the percentage of IgG bound to the antigen following treatment with denaturing agents (Ross et al., 2011).
Seroconversion remains a reliable means of diagnosing primary CMV infection but usually practical only for closely monitored patients such as transplant recipients, for whom pre and post infection sera are readily available (O’Neill et al., 1988 and Pass et al., 1983).
This is highly specific for the diagnosis of CMV infection (Razonable and Humar, 2013). Culture can be performed using the conventional plaque assay or the more rapid shell vial centrifugation culture system (Razonable et al., 2002). Isolation of CMV from most clinical samples (other than urine, saliva, and stool) is highly predictive of the diagnosis of CMV disease or the risk of progression from CMV infection into clinical illness (Razonable et al., 2002).
In contrast, the use of urine, saliva, and stool samples for CMV culture is of limited clinical utility because viral shedding may be detected in these specimens in CMV-seropositive patients even in the absence of clinical illness (Razonable and Humar, 2013).
For CMV-seronegative patients (seen most commonly in pediatric age groups), however, the isolation of CMV in urine (and other samples) may be clinically relevant, since it is suggestive of active primary infection (instead of shedding) (Razonable and Humar, 2013).
The major drawbacks to viral culture are its low to modest sensitivity and long turnaround time (Razonable et al., 2002). Accordingly, the clinical use of viral culture is minimal in the contemporary era, when molecular assays are most commonly used in the clinical setting (Razonable et al., 2002). The remaining major clinical use of viral culture is in the diagnosis of CMV infection by use of samples that have not been validated or optimized for molecular testing (Razonable and Humar, 2013). Viral culture may also be required when phenotypic antiviral drug resistance testing is needed, although advances in molecular genotypic assays have emerged for detecting antiviral drug resistance (Hakki and Chou, 2011).
CMV antigen detection in the blood is the most commonly used phenotypic method for the rapid and sensitive diagnosis of CMV infection (Razonable and Humar, 2013). CMV antigenemia assay uses monoclonal antibodies to detect the CMV pp65 antigen that is expressed in CMV-infected leukocytes during the early phase of the CMV replication process (Razonable et al., 2002). The result of the test is reported as the number of positive cells per total number of cells counted (Razonable and Humar, 2013).
Because pp65 is secreted during viral replication, its detection in peripheral blood leukocytes generally signifies active CMV infection. The CMV antigen assay is a rapid and easy test to perform and has a higher sensitivity than that of virus culture (Razonable and Hayden, 2013). It is able to detect CMV infection earlier than virus culture, with some studies reporting the detection of antigenemia an average of 5 to 14 days before the onset of CMV disease (Razonable and Hayden, 2013).
Thus, it can be used to detect early CMV replication and to guide the initiation of preemptive therapy (Singh, 2001). In general, the degree of pp65 antigenemia correlates with the risk of subsequent CMV disease. However, there is a lack of consensus as to the threshold of pp65-positive cells that should trigger the initiation of antiviral therapy (Razonable and Hayden, 2013).
In some studies, the sensitivity of pp65 antigenemia testing for the diagnosis of CMV infection was comparable to that of CMV NAT by PCR (Garrigue et al., 2006). One of these studies reported a strong correlation between pp65 antigenemia and CMV PCR performed on whole-blood specimens (Garrigue et al., 2006). Other studies, however, have reported a significantly lower sensitivity of antigenemia testing than those of molecular tests (Pang et al., 2009). Moreover, the plasma PCR assay detected CMV infection 12 days earlier than the antigenemia test (Hadaya et al., 2003).
The disadvantages of CMV antigenemia testing are its labor-intensive and manual nature. The interpretation of the test is subjective, and there is limited interlaboratory standardization of thresholds of positive cell counts to guide various clinical actions (Razonable et al., 2002). Blood samples being subjected to pp65 antigenemia testing should be processed rapidly (ideally within 6 h) to optimize sensitivity, since test results depend on the life span of leukocytes ex vivo. Delays in the processing of a sample for longer than 24 h may lead to a significant decrease in the number of detectable pp65-positive cells in the blood (Razonable and Hayden, 2013).
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