Description
Arthropod vectors, including mosquitoes, ticks, fleas, and mites, that feed on vertebrate hosts can spread bacteria, protozoa, and viruses during feeding to their susceptible host, resulting in a variety of infections and diseases. Arboviruses (arthropod-borne viruses) include Zika virus, West Nile virus (WNV), chikungunya virus, dengue virus (DENV), yellow fever virus (YFV), and Colorado tick fever virus (CTF) to name a few. Malaria and babesiosis are both conditions caused by arthropod-borne protozoan parasites, Plasmodium and Babesia, respectively. Conditions caused by arthropod-borne bacteria include rickettsial diseases, ehrlichiosis, anaplasmosis, and Lyme disease, as well as other Borrelia-associated disorders (Calisher, 1994; CDC, 2022a). Isolation, identification, and characterization of these various infections depend on the causative agent. Identification methods may include culture testing, microscopy, and staining techniques; moreover, molecular testing, such as nucleic acid amplification testing (NAAT), and serologic testing, including immunofluorescence antibody assays and enzyme-linked immunosorbent assays (ELISA), can be used for laboratory diagnosis (Miller et al., 2018).

For Lyme disease and testing for Borrelia burgdorferi, please see CAM 159 Lyme Disease.

Policy 
Application of coverage criteria is dependent upon an individual’s benefit coverage at the time of the request. 

1. For individuals suspected of having babesiosis (see Note 1), the use of a Giemsa- or Wright-stain of a blood smear or NAAT is considered MEDICALLY NECESSARY.
2. For individuals suspected of having babesiosis (see Note 1), the use of either an IgG or IgM indirect immunofluorescence antibody (IFA) assay for Babesia is considered NOT MEDICALLY NECESSARY.
3. For individuals suspected of having chikungunya virus (see Note 2), the use of viral culture for diagnosis, NAAT for the presence of chikungunya in a serum sample, or IFA assay for IgM antibodies during both the acute and convalescent phases is considered MEDICALLY NECESSARY.
4. For individuals suspected of having Colorado tick fever (CTF) (see Note 3), the use of virus-specific IFA-stained blood smears or IFA for CTF-specific antibodies is considered MEDICALLY NECESSARY.
5. For the detection of dengue virus (DENV), the use of NAAT, IgM antibody capture ELISA (MAC-ELISA), or NS1 ELISA, as well as a confirmatory plaque reduction neutralization test for DENV, is considered MEDICALLY NECESSARY in the following individuals:
   1. For individuals suspected of having DENV (see Note 4).
   2. For non-pregnant individuals who are symptomatic for Zika virus infection (see Note 5). 
6. For individuals suspected of having DENV (see Note 4), the use of IgG ELISA or hemagglutination testing is considered NOT MEDICALLY NECESSARY.
7. For individuals suspected of having ehrlichiosis and/or anaplasmosis (see Note 6), the use of NAAT of whole blood, IFA assay for IgG antibodies, or microscopy for morulae detection is considered MEDICALLY NECESSARY.
8. For individuals suspected of having ehrlichiosis and/or anaplasmosis (see Note 6), the use of an IFA assay for IgM antibodies or standard blood culture is considered NOT MEDICALLY NECESSARY.
9. For individuals suspected of having malaria (see Note 7), the use of a rapid immunochromatographic diagnostic test or smear microscopy to diagnose malaria, determine the species of Plasmodium, identify the parasitic life-cycle stage, and/or quantify the parasitemia (can be repeated up to three times within three days if initial microscopy is negative in suspected cases of malaria) is considered MEDICALLY NECESSARY.
10. For individuals suspected of having malaria (see Note 7), the use of NAAT or IFA for Plasmodium antibodies is considered NOT MEDICALLY NECESSARY.
11. For individuals suspected of having a rickettsial disease (see Note 8), the use of an IFA assay for IgG antibodies (limited to two units) is considered MEDICALLY NECESSARY.
12. For individuals suspected of having a rickettsial disease (see Note 8), the use of standard blood culture, nucleic acid amplification testing (NAAT), or IFA assay for IgM antibodies is considered NOT MEDICALLY NECESSARY.
13. For individuals suspected of having a tick-borne relapsing fever (TBRF) (see Note 9), the use of dark-field microscopy of a peripheral blood smear, microscopy of a Wright- or Giemsa-stained blood smear, PCR testing, or serologic assays to detect Borrelia specific IgG antibodies is considered MEDICALLY NECESSARY.
14. For individuals suspected of having a TBRF (see Note 9), the use of an IFA assay for IgM for Borrelia or culture testing for Borrelia is considered NOT MEDICALLY NECESSARY.
15. For individuals suspected of having West Nile virus (WNV) (see Note 10), the use of IFA for WNV-specific IgM antibodies in either serum or CSF and a confirmatory plaque reduction neutralization test for WNV is considered MEDICALLY NECESSARY.
16. For individuals suspected of having WNV (see Note 10), the use of NAAT for WNV or IFA for WNV-specific IgG antibodies in either serum or CSF is considered NOT MEDICALLY NECESSARY.
17. For individuals suspected of having yellow fever virus (YFV) (see Note 11), the use of NAAT for YFV or serologic assays to detect virus-specific IgM and IgG antibodies, as well as a confirmatory plaque reduction neutralization test for YFV, is considered MEDICALLY NECESSARY
18. For the detection of Zika virus, the use of NAAT is considered MEDICALLY NECESSARY in the following individuals:
    1. Up to 12 weeks after the onset of symptom for symptomatic (see Note 5) pregnant individuals who have either recently traveled to areas with a risk of Zika (see Note 12) or who have had sex with someone who either lives in or has recently traveled to areas with a risk of Zika (see Note 12).
    2. For infants born from individuals who, during pregnancy, tested positive for Zika virus. 
    3. For infants born with signs and symptoms of congenital Zika syndrome (see Note 13) and who have a birthing parent who, during pregnancy, traveled to an area with a risk of Zika (see Note 12). 
19. For pregnant individuals who have a fetus with prenatal ultrasound findings consistent with congenital Zika virus infection (see Note 13), Zika virus NAAT (maternal serum and maternal urine) and Zika virus IgM testing (maternal serum), as well as a confirmatory plaque reduction neutralization test for Zika, is considered MEDICALLY NECESSARY.  
20. For non-pregnant individuals symptomatic for Zika virus infection (see Note 5), NAAT and/or IgM testing for Zika detection is considered NOT MEDICALLY NECESSARY.
21. For asymptomatic individuals, testing for babesiosis, chikungunya virus, CTF, DENV, ehrlichiosis and/or anaplasmosis, malaria, rickettsial disease, TBRF, WNV, YFV, or Zika virus during a general exam without abnormal findings is considered NOT MEDICALLY NECESSARY.

NOTES:

Note 1: Typical signs and symptoms of babesiosis can include hemolytic anemia, splenomegaly, hepatomegaly, jaundice, and nonspecific flu-like symptoms such as fever, chills, body aches, weakness, and fatigue (CDC, 2019a).

Note 2: Typical signs and symptoms of chikungunya include high fever (> 102^◦F or 39^◦C), joint pains (usually multiple joints, bilateral, and symmetric), headache, myalgia, arthritis, conjunctivitis, nausea, vomiting, and maculopapular rash (Staples et al., 2020).

Note 3: Typical signs and symptoms of CTF can include fever, chills, headache, myalgia, malaise, sore throat, vomiting, abdominal pain, and maculopapular or petechial rash (CDC, 2023a).

Note 4: Typical signs and symptoms of dengue can include fever, headache, retro-orbital eye pain, myalgia, arthralgia, erythematous maculopapular rash, petechiae, leukopenia, and nausea and/or vomiting (CDC, 2019b).

Note 5: Typical signs and symptoms of Zika virus infection can include fever, rash, headache, joint pain, conjunctivitis (red eyes), and muscle pain (CDC, 2019d).

Note 6: Typical signs and symptoms of ehrlichiosis and/or anaplasmosis usually begin 5 – 14 days after an infected tick bite, and they include fever, headache, malaise, myalgia, and shaking chills. Ehrlichiosis can also present with gastrointestinal issues, including nausea, vomiting, and diarrhea (Biggs et al., 2016).

Note 7: Typical signs and symptoms of malaria can include fever, influenza-like symptoms (e.g., chills, headache, body aches), anemia, jaundice, seizures, mental confusion, kidney failure, and acute respiratory distress syndrome (Arguin & Tan, 2019).

Note 8: Typical signs and symptoms of rickettsial diseases (including Rocky Mountain spotted fever, Rickettsia parkeri rickettsiosis, Rickettsia species 364D rickettsiosis, Rickettsia spp (mild spotted fever), and R. akari (rickettsialpox)) usually begin 3 – 12 days after initial bite and can include fever, headache, chills, malaise, myalgia, nausea, vomiting, abdominal pain, photophobia, anorexia, and skin rash. Rickettsia species 364d rickettsiosis can also present with an ulcerative lesion with regional lymphadenopathy (Biggs et al., 2016).

Note 9: Typical signs and symptoms of tick-borne relapsing fever (caused by Borrelia hermsii, B. mazzottii, B. miyamotoi, B. parkeri, or B. turicatae) include recurring febrile episodes that last approximately 3 days separated by approximately 7 days. Nonspecific symptoms that occur in at least 50% of cases include headache, myalgia, chills, nausea, arthralgia, and vomiting (CDC, 2022e).

Note10: Typical signs and symptoms of WNV include headache, myalgia, arthralgia, gastrointestinal symptoms, and maculopapular rash. Less than 1% of infected individuals develop neuroinvasive WNV with symptoms of meningitis, encephalitis, or acute flaccid paralysis (Nasci et al., 2013).

Note 11: Typical signs and symptoms of yellow fever include symptoms of the toxic form of the disease (jaundice, hemorrhagic symptoms, and multisystem organ failure), as well as nonspecific influenza symptoms (fever, chills, headache, backache, myalgia, prostration, nausea, and vomiting in initial illness) (Gershman & Staples, 2021).

Note 12: The CDC provides information on the risk of Zika in areas in the United States (https://www.cdc.gov/zika/geo/index.html) and outside of the United States and its territories (https://wwwnc.cdc.gov/travel/page/zika-information). 

Note 13: Typical signs and symptoms of congenital Zika syndrome can include microcephaly, problems with brain development, feeding problems (e.g., difficulty swallowing), hearing loss, seizures, vision problems, decreased joint movement (i.e., contractures), and stiff muscles (making it difficult to move) (CDC, 2022b).

Rationale
Hematophagous arthropods, such as mosquitoes, ticks, fleas, and mites, can spread opportunistic bacteria, protozoa, and viruses to host organisms when feeding. Numerous outbreaks of arthropod-borne disease have been documented, including plague, an acute febrile disease caused by Yersinia pestis through the bite of infected fleas, which resulted in more than 50 million deaths in Europe alone during the “Black Death” outbreak. More than 3000 cases of plague were reported to the World Health Organization (WHO) between 2010 and 2015 with 584 deaths. Today, most cases of plague occur in the Democratic Republic of Congo, Madagascar, and Peru (WHO, 2017).

The Centers for Disease Control and Prevention (CDC) reported a large increase in the number of vector-borne diseases within the United States and its territories between 2004 – 2016. More than 640,000 cases were reported during that time; in fact, infections of tick-borne bacteria and protozoa more than doubled from 2004 to 2016. “In the United States, 16 vectorborne diseases are reportable to state and territorial health departments, which are encouraged to report them to the National Notifiable Disease Surveillance System (NNDSS). Among the diseases on the list that are caused by indigenous pathogens are Lyme disease (Borrelia burgdorferi); West Nile, dengue, and Zika virus diseases; plague (Yersinia pestis); and spotted fever rickettsioses (e.g., Rickettsia rickettsii). Malaria and yellow fever are no longer transmitted in the United States but have the potential to be reintroduced” (Rosenberg et al., 2018). New vector-borne infections are emerging; for example, two unknown, life-threatening RNA viruses spread by ticks have been identified in the U.S. since 2004. Although both tick- and mosquito-borne diseases are increasing across the U.S., the CDC reports that these two vectors are showing different trends. The mosquito-borne diseases are characterized by epidemics; for example, West Nile Virus is essentially limited to the continental U.S. but has spread rapidly since its introduction to New York in 1999, whereas chikungunya and dengue primarily occur within the U.S. territories. On the other hand, the tick-borne disease increase occurs in the continental U.S. and has experienced a gradual, steady rate increase with Lyme disease comprising 82% of all tick-borne diseases (Rosenberg et al., 2018). Figure 1 and 2 below, taken from Rosenberg et al. (2018), show the reported cases of tickborne and mosquito-borne disease in the United States from 2004 – 2016.
6.

Rickettsial infections
Rocky Mountain spotted fever (RMSF) is the most common rickettsial infection in the U.S. with 6,248 cases reported to the CDC alone in 2017 (CDC, 2022d). RMSF is caused by Rickettsia rickettsia, spread in the U.S. predominantly by Dermacentor variabilis (the American dog tick) and D. andersoni (the Rocky Mountain wood tick), and can be found throughout North America as well as parts of South America. The Council for State and Territorial Epidemiologists combined RMSF with other rickettsial diseases into the more broad “spotted fever rickettsiosis” designation in 2010 (CDC, 2022d). Besides the obligatory tick bite, typical symptoms of RMSF include fever, headache, and rash with the characteristic rash occurring in approximately 88% to 90% of patients within three to five days of illness. If left untreated, RMSF can be fatal but can easily be treated with antimicrobial therapy upon timely diagnosis. Definitive diagnosis of RMSF cannot usually be made via culture because Rickettsia cannot be grown in cell-free culture media; they are obligate intracellular bacteria requiring living host cells. RMSF diagnosis can be made via either skin biopsy prior to treatment with antibiotics or through serologic testing using indirect immunofluorescence assays (IFAs). Immunoglobulin G (Biggs et al.) antibodies are more specific than immunoglobulin M (IgM) antibodies since the latter can give false-positive results due to cross-reactivity with other bacterial pathogens. A drawback of IFA is that usually it is unreliable for the first five days of infection until antibody levels are high enough for detection. The CDC and major clinical labs do offer a polymerase chain reaction (PCR)-based assay for RMSF (Sexton & McClain, 2023a).

Since 2001, thirteen more human rickettsiae belonging to the spotted fever group (SFG) have been identified. All SFGs can cause fever, headache, and myalgia and are arthropod-borne (primarily ticks and mites). Most patients with an SFG display a rash and/or a localized eschar. Rickettsialpox, caused by R. akari, is transmitted from the bite of a house mouse mite, usually after mouse extermination programs result in a decrease of the mite’s food supply. Rickettsialpox is typically a relatively mild disease that can resolve itself without treatment within three weeks, but treatment hastens improvement. Rickettsiosis can also be due to infection with R. parkeri, R. amblyommii, and Rickettsia species 364D (also called R. philipii). Isolation of SFG rickettsiae is rare in clinical practice due to the difficulty of obtaining culture; consequently, serology, immunologic detection from tissue, and PCR are more often used for diagnosis. Microimmunofluorescent (MIF) antibody tests, enzyme-linked immunosorbent assays (ELISAs), and Western blot immunoassays can be used to detect convalescent IgG and IgM antibodies, but these methods can only be used at least 10 – 14 days after the onset of illness when antibody concentrations are high enough for detection. McQuiston et al. (2014) concluded that the “use of IgM antibodies should be reconsidered as a basis for diagnosis and public health reporting of RMSF and other spotted fever group rickettsia in the United States” in one small study; the study demonstrated that IgM findings often resulted in false positives for Rock Mountain Spotted Fever and questioned the value of IgM testing (McQuiston et al., 2014).PCR is a very specific technique. PCR using tissue samples has higher specificity than whole blood PCR. Immunologic detection from a tissue biopsy requires the use of special laboratory equipment so it is not as frequently used as either the serologic or PCR detection methods (Sexton & McClain, 2023c). 

Ehrlichiosis and Anaplasmosis
Human ehrlichiosis was first reported in 1986, and the causative agent for human granulocytic anaplasmosis, Anaplasma phagocytophilum, was identified in 1994. Both ehrlichiosis and anaplasmosis are transmitted from the bite of infected ticks and have similar clinical and laboratory manifestations. Ehrlichiosis can be caused by Ehrlichia chaffeensis, E. ewingii, and E. muris. Typically, patients have a fever within an incubation period of one to two weeks. Other symptoms can include malaise, myalgia, headache, chills, gastrointestinal distress, and cough. Both leukopenia and thrombocytopenia can occur. Diagnosis via culture is extremely difficult. “Until 1995, only two isolates of E. chaffeensis had been recovered from humans; in both cases, this process required over 30 days of cultivation. The isolation of A. phagocytophilum from three additional patients has been accomplished using a cell culture system derived from human promyelocytic leukemia cells (Sexton & McClain, 2023b). IFA testing for bacteria-specific antibodies is the most common method for diagnosing ehrlichiosis and anaplasmosis, but similar to rickettsiae, ELISA, PCR, and immunochemical tissue staining can be used as well. Unlike rickettsiosis, ehrlichiosis and anaplasmosis can also be detected by the presence of characteristic intraleukocytic morulae in a peripheral blood smear or buffy coat smear (Sexton & McClain, 2022).   

Borrelia Infections
Besides Lyme disease, caused by Borrelia burgdorferi, Borrelia can cause relapsing fever. Tick-borne relapsing fever (TBRF) in North America is primarily caused by B. hermsii, B. turicatae, B. parkeri, B. miyamotoi, and B. mazzottii, and louse-borne relapsing fever (LBRF) is an infection caused by B. recurrentis (Barbour, 2023; Miller et al., 2018). The characteristic feature of these infections is the relapsing fever due to cyclical spirochetemia caused by antigenic variation of the spirochetes. Each bout of fever lasts 3 to 12 days with temperatures ranged from 39^◦C to 43^◦C (102.2^◦F to 109.4^◦F). Visual analysis by Giemsa or Wright staining blood smears taken during a febrile episode is common practice. PCR can also be used on a variety of samples, including cerebrospinal fluid (CSF), blood, tissue, or even culture medium. According to the CDC, “a change in serology results from negative to positive, or the development of an IgG response in the convalescent sample, is supportive of a TBRF diagnosis” (CDC, 2022c). One exception is using antibodies to the GlpQ protein characteristic of these Borrelia species but not to B. burgdorferi (Lyme disease) (Barbour, 2020).

Protozoa infections
Babesiosis is due to primarily Babesia microti in the U.S, but B. divergens and B. venatorum are the primary causative agents of babesiosis in Europe and China, respectively. The incubation period of Babesia depends on the mode of transfection: 1 – 4 weeks following a tick bite; the incubation period after transfusion of contaminated blood products usually or three to seven weeks but ranges from one week to six months. The most common symptoms of infection include a fever, fatigue, malaise, chills, sweats, headache, and myalgia. Immunocompromised individuals can develop relapsing babesiosis due to an absent or impaired production of antibodies with approximately 20% mortality rate for patients who develop relapsing babesiosis. Most patients with babesiosis are also co-infected with other tick-borne bacterial pathogens. “Preferred tools for diagnosis of babesiosis include blood smear for identification of Babesia organisms and polymerase chain reaction (PCR) for detection of Babesia DNA. Serology can be a useful adjunct to blood smear and PCR” (Krause & Vannier, 2023). Serology is not ideal in diagnosing an acute infection since antibody concentrations remain elevated post-recovery.

Plasmodium falciparum, P. vivax, and P. ovale are responsible for malaria. They are spread by the bite of an Anopheles mosquito where their sporozoites infect the liver within one to two hours. Within the hepatocyte, they form merozoites. Upon rupturing into the bloodstream, the merozoites infect red blood cells for trophozoite formation, causing the erythrocytic stage of the life cycle where additional merozoites are released. During this stage of the cycle, the symptoms of malaria, including fever, occur. This process usually takes 12 to 35 days, but clinical manifestations can be delayed in individuals with partial immunity or those who are taking ineffective prophylaxis. Other initial symptoms can include irregular heartbeat, cough, anorexia, gastrointestinal distress, sweating, chills, malaise, arthralgia, and myalgia. Malaria, if left untreated, can also include acidosis, hypoglycemia, severe anemia, renal and hepatic impairment, edema, and death (Cohee & Seydel, 2023). Parasite-based diagnosis may include microscopic examination of blood smears, which can often identify the species of Plasmodium as well as the parasite density, and antigen-based tests. Rapid diagnostic testing (RDT) of the antigens using immunochromatographic methods is available, but the accuracy of the RDT can vary considerably. NAATs can also be used to identify a malarial infection, and NAATs “are typically used as a gold standard in efficacy studies for antimalarial drugs, vaccines, and evaluation of other diagnostic agents” with a “theoretical limit of detection for PCR … estimated at 0.02 to 1 parasite/microL” (Hopkins, 2023). The Mayo Clinic Laboratories indicates that “PCR is an alternative method of malaria diagnosis that allows for sensitive and specific detection of Plasmodium species DNA from peripheral blood. PCR may be more sensitive than conventional microscopy in very low parasitemias, and is more specific for species identification … Malaria PCR can be used in conjunction with traditional blood film or Babesia PCR when the clinical or morphologic differential includes both babesiosis and malaria” "Test ID: LCMAL Malaria, Molecular Detection, PCR, Varies" (2023).

Viral infections
Examples of arthropod-borne viruses (arboviruses) include West Nile virus (WNV), dengue, yellow fever virus (YFV), chikungunya, and Colorado tick fever virus. In the United States, WNV is the most common arbovirus reported to the CDC. In 2016, 96% of the reported 2,240 cases of domestic arboviruses were WNV with 61% of the WNV cases reported being neuroinvasive. Neuroinvasive WNV includes meningitis, encephalitis, and acute flaccid paralysis (Burakoff et al., 2018). In general, most infected individuals are asymptomatic with only 20 – 40% of infected patients showing any characteristic symptoms of WNV, including fever, headache, malaise, myalgia, anorexia, and rash. Diagnosis of WNV of a symptomatic individual usually occurs with a WNV IgM antibody capture ELISA (MAC-ELISA) assay. A patient with symptoms of a neurologic infection does require a lumbar puncture. Confirmatory testing can include a plaque reduction neutralization test (PRNT). PCR testing is primarily used with immunocompromised patients who have delayed or absent antibody production, patients with a history of prior flavivirus infections, and blood donors who may be asymptomatic (Lyle R Petersen, 2023).

Dengue virus (DENV) infection is a result of being bitten by an infected Aedes aegypti or A. albopictus mosquito. Four distinct DENV types of Flavivirus are known: DENV-1, DENV-2, DENV-3, and DENV-4. DENV is endemic throughout much of the tropical regions of the world, but the only region of the U.S. endemic for DENV is Puerto Rico. The last major outbreak occurred in Puerto Rico in 2010 where 26,766 cases of suspected DENV were reported and 47% of all laboratory tested specimen were positive (CDC, 2019b, 2023c). “Dengue fever … is an acute febrile illness defined by the presence of fever and two or more of the following but not meeting the case definition of dengue hemorrhagic fever: headache, retro-orbital or ocular pain, myalgia and/or bone pain, arthralgia, rash, hemorrhagic manifestations … [and] leukopenia. The cardinal feature of dengue hemorrhagic fever is plasma leakage due to increased vascular permeability as evidenced by hemoconcentration (≥ 20 percent rise in hematocrit above baseline), pleural effusion, or ascites. DHF [dengue hemorrhagic fever] is also characterized by fever, thrombocytopenia, and hemorrhagic manifestations …. (Thomas et al., 2023).” Laboratory diagnostic testing includes direct detection of viral components in serum or indirect serologic assays. “Detection of viral nucleic acid or viral antigen has high specificity but is more labor intensive and costly; serology has lower specificity but is more accessible and less costly” (Thomas et al., 2023). Culture testing as a diagnostic tool usually is time-prohibitive.

Zika virus is a mosquito-borne illness discovered in Uganda in 1947 but has since spread across Asia and to the Americas. Zika infection has been tied to several birth defects. The first human cases of Zika were detected in 1952. Prior to 2007, at least 14 cases of Zika had been documented. Symptoms of Zika are similar to those of many other diseases; therefore, many cases may not have been recognized (CDC, 2019c). The most common symptoms of Zika are fever, rash, joint pain, and conjunctivitis (CDC, 2019c). The illness is usually mild with symptoms beginning 2 – 7 days after being bitten by an infected mosquito, lasting for several days to a week. Most individuals infected with Zika virus are unaware of the infection, as only a maximum of 25% of people infected will exhibit symptoms(CDC, 2019c; LeBeaud, 2021). Diagnosis of the Zika virus is definitively established through reverse-transcription polymerase chain reaction (RT-PCR) for Zika virus RNA in all symptomatic patients. Aside from pregnant individuals who have traveled to an at risk area, asymptomatic patients are typically not tested (LeBeaud, 2021).

Colorado tick fever virus (CTFV) is a Reoviridae transmitted primarily by the Rocky Mountain wood tick (Dermacentor andersoni) in the western U.S. and Canada. Transmission of CTFV has also been reported in blood transfusions. The incubation period can last up to 14 days, and symptoms include fever, headache, chills, myalgia, leukopenia, and prostration. Only 15% of symptomatic patients demonstrate a rash. Serologic tests are usually not helpful until at least 10 – 14 days for antibody production whereas real-time PCR (RT-PCR) can be used on the first day of symptoms (L. R. Petersen, 2023).

Yellow fever, occurring primarily in sub-Saharan Africa and South America, is a flavivirus spread by mosquitoes that causes hemorrhagic fever with a high fatality rate. An outbreak in Brazil in January – March 2018 resulted in 4 of 10 patients infected with YFV dying. None of those showing symptoms had been vaccinated against YFV. Yellow fever causes hemorrhagic diathesis due to decreased synthesis of vitamin K-dependent coagulation factors as well as hepatic dysfunction, renal failure, and coagulopathy. Yellow fever diagnosis is typically made by a serologic test using an ELISA-IgM assay; however, this assay does cross-react with other flaviviruses and with the YFV vaccination. Rapid diagnostic testing using either PCR or immunoassay is available. Viral isolation and culture can be performed, but it requires inoculation of mosquitoes or mammalian cell culture. Tissue biopsy, such as liver, cannot be performed on the living patient due to possible fatal hemorrhaging; biopsy would be performed during the post-mortem workup (Wilder-Smith, 2023).

Chikungunya virus, endemic in many tropical and subtropical regions of the world, is transmitted by the mosquitoes Aedes aegypti and Aedes albopictus. Within the U.S., chikungunya is prevalent in Puerto Rico where approximately 25% of blood donors were seropositive; it has also been reported in Florida. Both dengue and Zika are transmitted by the same vectors, so these viruses often co-circulate geographically Chikungunya can cause acute febrile polyarthralgia and arthritis. The predominant testing method for diagnosis of chikungunya is the detection of viral RNA via either RT-PCR or virus serology using either ELISA or IFA. Viral culture is typically not used as a diagnostic tool but is used for epidemiologic research (Wilson & Lenschow, 2023).

Types of Testing

Analytical Validity 
The use of antibodies to detect and diagnose arthropod-associated infections and diseases is a common practice. Johnson et al. (2000) first reported the use of monoclonal antibody-based capture ELISA testing for a variety of alphaviruses, including chikungunya, flaviviruses, including dengue and yellow fever, and bunyaviruses. The researchers concluded, “IgG ELISA results correlated with those of the standard plaque-reduction neutralization assays. As expected, some test cross-reactivity was encountered within the individual genera, and tests were interpreted within the context of these reactions. The tests were standardized for laboratory diagnosis of arboviral infections, with the intent that they be used in tandem with the corresponding IgM antibody-capture ELISAs” (Johnson et al., 2000). Kalish and associates also demonstrated that IgG and/or IgM antibody responses can still occur up to 20 years post-infection; consequently, a rise in antibody titer does not necessarily indicate a current, acute infection (Kalish et al., 2001).

Granger and Theel (2019) published an evaluation of two enzyme-linked immunosorbent assays and a rapid immunochromatographic assay for the detection of IgM antibodies to Zika virus. This article states that five serological assays have been approved by the FDA in an emergency use situation and include the Chembio DPP Zika IgM system (a rapid immunochromatographic assay), the InBios ZIKV Detect 2.0 IgM antibody capture enzyme-linked immunosorbent assay, and the InBios ZIKV Detect MAC-ELISA. These three serologic assays were evaluated, using 72 samples, based on the identification of neutralizing antibodies to Zika virus, dengue virus, or West Nile virus. “The Chembio DPP Zika ICA and InBios ZIKV 2.0 MAC-ELISA showed 95% specificity in 22 ZIKV/DENV-seronegative specimens and in 13 samples positive for NAbs to non-ZIKV flaviviruses. Comparatively, the InBios ZIKV MAC-ELISA was “presumptive” or “possible Zika positive” in 8 of 12 WNV or DENV PRNT-positive samples and in 12 of 22 PRNT-seronegative sera (Granger & Theel, 2019).” The authors conclude that by replacing the InBios ZIKV MAC-ELISA with the InBios ZIKV 2.0 MAC-ELISA, testing burden will be minimized on laboratories performing PRNT for the identification of neutralizing antibodies.

Leski et al. (2020) performed a 2020 study published in Malaria Journal that compared traditional diagnostic methods such as rapid diagnostic tests (RDTs) and DNA-based methods to polymerase chain reaction (PCR). The results indicated consistency with “previous observations that PCR-based tests have a significantly higher sensitivity when compared with both microscopy and RDTs” Leski et al. (2020).

Mathison and Pritt (2017) reviewed current standards for malaria testing and the most used methods for laboratory diagnosis. The most common tests “are microscopic examination of stained blood flims and detection of parasite antigen or nucleic acid … Rapid antigen detection methods and molecular amplification tests are also increasingly employed for malaria diagnosis and are useful adjunctive tests.” According to the algorithm developed in “Update on Malaria Diagnostics and Test Utilization,” NAAT tests are one of three tests recommended for use if malaria is suspected based on clinical findings and exposure history (Mathison & Pritt, 2017).

Kim et al. (2018) had also developed a rapid diagnostic test (RDT) for detecting IgG/IgM antibodies against Zika virus using “monoclonal antibodies to the envelope (E) and non-structural protein (NS1).” The diagnostic accuracy of this kit was “fairly high; sensitivity and specificity for IgG was 99.0 and 99.3%, respectively, while for IgM it was 96.7 and 98.7%, respectively.” However, there were cross reactions with the dengue virus evaluated using anti-Dengue Mixed Titer Performance Panel (PVD201), “in which the Zika RDT showed cross-reactions with [dengue virus] in 16.7% and 5.6% in IgG and IgM, respectively.” This research could potentially enable the rapid diagnostic test to be preferable to the traditional RT-PCR in endemic areas (Kim et al., 2018).

Clinical Utility and Validity
In 2013, Kato and colleagues tested the sensitivity of two different RT-PCR-based assays for Rickettsia — PanR8, an assay that tests for Rickettsia in general, and RRi6, an assay specific for R. rickettsii. Both of these methods were more sensitive in testing for Rickettsia than the nested PCR method of the CDC; moreover, both of these methods are faster than the nested PCR method (1 hr versus 1 – 2 days, respectively) (Kato et al., 2013). These results were corroborated in 2014 by Denison and colleagues. They used a multiplex PCR assay to correctly identify all cell controls for R. rickettsii, R. parkeri, and R. akari; moreover, no false-positive results were reported using this methodology. “This multiplex real-time PCR demonstrates greater sensitivity than nested PCR assays in FFPE [formalin-fixed, paraffin-embedded] tissues and provides an effective method to specifically identify cases of Rocky Mountain spotted fever, rickettsialpox, and R. parkeri rickettsiosis by using skin biopsy specimens” (Denison et al., 2014).

The FDA has approved the use of the BinaxNOW malaria test for screening and diagnosing malaria. Even though this testing method is considerably faster than other methods (as low as 1.1 – 1.7 hours complete turnaround time (Ota-Sullivan & Blecker-Shelly, 2013), the use of BinaxNOW in non-endemic areas is a point of controversy due to relatively low sensitivity (84.2%) and for misclassifying Plasmodium falciparum malaria as non-falciparum (Dimaio et al., 2012). Moreover, it has been reported that Salmonella typhi can give a false-positive for malaria using the BinaxNOW test (Meatherall et al., 2014). 

van Bergen et al. (2021) evaluated a novel real-time PCR assay for clinical validity. The authors used reference samples, patient samples, and synthetic controls. The analytical performance details of the MC004 assay were considered: “analytical specificity, limit of detection, the ability to detect mixed infections, and the potential to determine the level of parasitaemia of P. falciparum, including assessment of within-run and between-run precisions.” The authors reported “zero false positive or false negative results.” Regarding precision, “the within-run and between-run precisions were less than 20% CV at the tested parasitaemia levels of 0.09%, 0.16%, 2.15% and 27.27%.” Based on these results, the authors reported that “the entry of PCR-based techniques into malaria diagnostics has improved the sensitivity and specificity of the detection of Plasmodium infections … Based upon the analytical performance characteristics that were determined, the MC004 assay showed performance suitable for use in clinical settings, as well as epidemiological studies” (van Bergen et al., 2021).

Akoolo et al. (2017) compared qPCR results in the detection of Babesia infection against currently available non-NAAT tests (FISH and microscopy). Blood samples were analyzed from 192 patients. The researchers report that “Of 28 samples that were positive by FISH, 27 (96%) were also positive by qPCR indicating high congruency between nucleic acid-based tests. Interestingly, of 78 asymptomatic samples not tested by FISH, 22 were positive by our qPCR” (Akoolo et al., 2017). Overall, the qPCR method was found to have a sensitivity of 96.2% and a specificity of 70.5%. The authors conclude, “Robust qPCR using specific probes can be highly useful for efficient and appropriate diagnosis of babesiosis in patients in conjunction with conventional diagnostics, or as a stand-alone test, especially for donated blood screening” (Akoolo et al., 2017).
Reynolds et al. (2017) examined the 2016 United States Pregnancy Registry to estimate the proportion of birth defects of pregnant women exposed to Zika, and out of 972 pregnancies with laboratory evidence of a possible Zika infection, 51 had birth defects (5%). Of the 250 confirmed infections, 24 had birth defects. Similarly, Shiu et al. (2018) evaluated the screening results of the Zika virus in Miami-Dade County in Florida. Of 2327 women screened for Zika, 86 had laboratory evidence of infection, and 2 had congenital Zika “syndrome” (Zika-caused birth defects) (Shiu et al., 2018).

Centers for Disease Control and Prevention (CDC) 
Diagnosis and Management of Tickborne Rickettsial Diseases (Biggs et al., 2016): In 2016, the CDC released their guidelines and recommendations concerning Rickettsial diseases, including Rocky Mountain spotted fever, in the MMWR. The table below summarizes their recommended diagnostic tests for tickborne rickettsial diseases:

To summarize their recommendations, even though indirect immunofluorescence antibody assays (IFAs) are insensitive typically during the first week of an acute infection, they are the standard reference for tickborne rickettsial infections; in addition, a minimum of two tests are to be performed for a diagnosis. Usually, one sample is taken early after the initial symptoms are present, and a second sample is taken 2 – 4 weeks later. A minimum of a fourfold rise in antibody titer is required to confirm diagnosis. In cases of ehrlichiosis and anaplasmosis, during the first week, PCR amplification can be used on whole blood for diagnosis, but PCR has low sensitivity in Rocky Mountain spotted fever except in patients with severe disease. Morulae detection via either blood-smear or buffy-coat preparation microscopy can also be indicative of ehrlichiosis or anaplasmosis. However, “Rickettsiae cannot be isolated with standard blood culture techniques because they are obligate intracellular pathogens; specialized cell culture methods are required. Because of limitations in availability and facilities, culture is not often used as a routine confirmatory diagnostic method for tickborne rickettsial diseases” (Biggs et al., 2016).

Tick-borne relapsing fever (TBRF) (CDC, 2022e): In the U.S., TBRF can be caused by Borrelia hermsii, B. parkerii, and B. turicatae with B. hermsii being the most common causative agent. TBRF often presents with a relapsing nature (usually ~ 3 days per febrile episode followed by an afebrile period of approximately one week). Moreover, “Spirochetemia (spirochetes in blood) in TBRF patients often reaches high concentrations (> 106 spirochetes/ml). Thus, microscopy is a useful diagnostic tool for TBRF. The diagnosis of TBRF may be based on direct microscopic observation of relapsing fever spirochetes using dark field microscopy or stained peripheral blood smears. Spirochetes are more readily detected by microscopy in symptomatic, untreated patients early in the course of infection. Other bacteria, such as Helicobacter, may appear morphologically similar, so it is important to consider clinical and geographical characteristics of the case when making a diagnosis of TBRF based on microscopy. Additional testing, such as serology or culture, is recommended.”

CDC acknowledges that “Serologic testing for TBRF is not standardized and results may vary by laboratory. Serum taken early in infection may be negative, so it is important to also obtain a serum sample during the convalescent period (at least 21 days after symptom onset). A change in serology results from negative to positive, or the development of an IgG response in the convalescent sample, is supportive of a TBRF diagnosis. However, early antibiotic treatment may limit the antibody response. Patients with TBRF may have false-positive tests for Lyme disease because of the similarity of proteins between the causative organisms. A diagnosis of TBRF should be considered for patients with positive Lyme disease serology who have not been in areas endemic for Lyme disease. Incidental laboratory findings include normal to increased white blood cell count with a left shift towards immature cells, a mildly increased serum bilirubin level, mild to moderate thrombocytopenia, elevated erythrocyte sedimentation rate (ESR), and slightly prolonged prothrombin time (Reynolds et al.) and partial thromboplastin time (PTT)” (CDC, 2022e). 

Colorado Tick Fever (CTF) (CDC, 2023b): As of 2015, CTF was reportable in Arizona, Colorado, Montana, Oregon, Utah, and Wyoming. “Laboratory diagnosis of CTF is generally accomplished by testing of serum to detect viral RNA or virus-specific immunoglobulin (Ig) M and neutralizing antibodies. Antibody production can be delayed with CTF, so tests that measure antibodies may not be positive for 14 – 21 days after the onset of symptoms. RT-PCR (reverse-transcriptase polymerase chain reaction) is a more sensitive test early in the course of disease. CTF testing is available at some commercial and state health department laboratories and at CDC. Contact your state or local health department for assistance with diagnostic testing. They can help you determine if samples should be sent to the CDC Arbovirus Diagnostic Laboratory for further testing” (CDC, 2023b). 

Babesiosis (CDC, 2019a): According to the CDC website, the most recent update about babesiosis for health professionals is from 2012 (with revision in 2018). Diagnosis can be challenging due to the nonspecific clinical manifestations of the disease. “For acutely ill patients, the findings on routine laboratory testing frequently include hemolytic anemia and thrombocytopenia. Additional findings may include proteinuria, hemoglobinuria, and elevated levels of liver enzymes, blood urea nitrogen, and creatinine. If the diagnosis of babesiosis is being considered, manual (non-automated) review of blood smears should be requested explicitly. In symptomatic patients with acute infection, Babesia parasites typically can be detected by light-microscopic examination of blood smears, although multiple smears may need to be examined. Sometimes it can be difficult to distinguish between Babesia and Plasmodium (especially P. falciparum) parasites and even between parasites and artifacts (such as stain or platelet debris). Consider having a reference laboratory confirm the diagnosis — by blood-smear examination and, if indicated, by other means, such as molecular and/or serologic methods tailored to the setting/species” (CDC, 2019a). 

Malaria (Arguin & Tan, 2019): The CDC considers smear microscopy as the gold standard in diagnosing malaria since it can determine the species, identify the stage of parasitic life-cycle, and quantify the parasitemia. The CDC states, “Blood smear microscopy remains the most important method for malaria diagnosis. Microscopy can provide immediate information about the presence of parasites, allow quantification of the density of the infection, and allow determination of the species of the malaria parasite — all of which are necessary for providing the most appropriate treatment. Microscopy results should ideally be available within a few hours. These tests should be performed immediately when ordered by a health care provider. They should not be saved for the most qualified staff to perform or batched for convenience. In addition, these tests should not be sent out to reference laboratories with results available only days to weeks later. Assistance with speciation of malaria on smears is available from CDC” (Arguin & Tan, 2019). The CDC also notes that rapid diagnostic tests (RDTs) for malaria can detect malaria parasitic antigens. However, “RDTs offer a useful alternative to microscopy in situations where reliable microscopic diagnosis is not immediately available. Although RDTs can detect malaria antigens within minutes, they have several limitations. RDTs cannot distinguish between all of the Plasmodium species that affect humans, they may be less sensitive than expert microscopy or PCR for diagnosis, they cannot quantify parasitemia, and an RDT-positive test result may persist for days or weeks after an infection has been treated and cleared. Thus, RDTs are not useful for assessing response to therapy. Both positive and negative RDT results must always be confirmed by microscopy. Microscopy confirmation of the RDT result should occur as soon as possible because the information on the presence, density, and parasite species is critical for optimal management of malaria”(Arguin & Tan, 2019). Regarding PCR, the CDC states that “Although these tests are more sensitive than routine microscopy, results are not usually available as quickly as microscopy results, thus limiting the utility of this test for acute diagnosis and initial clinical management. Use of PCR testing is encouraged to confirm the species of malaria parasite and detect mixed infections” (Arguin & Tan, 2019).
The CDC also provided an update to malaria diagnosis in 2018. Although microscopy remained the “gold standard” for confirmation of malaria, other tests such as RDTs and PCR-based tests remained useful in certain situations (namely if microscopy is unavailable). PCR is considered most useful for “confirming the species of malarial parasite after the diagnosis has been established by either smear microscopy or RDT.” Finally, the CDC recommends that all cases of malaria be evaluated for drug resistance, typically through molecular characterization (PCR, gene sequencing) or in vitro tests (CDC, 2018b). 

Chikungunya (Staples et al., 2017, 2020): In the CDC Yellow Book, concerning the chikungunya virus, they recommend that “the differential diagnosis of chikungunya virus infection depends on the clinical signs and symptoms as well as where the person was suspected of being infected.” The other diseases to consider include: Zika, malaria, leptospirosis, parvovirus, group A Streptococcus, rubella, measles, dengue, enterovirus, adenovirus, alphavirus infections, post-infectious arthritis, and rheumatic conditions. Laboratory diagnosis is done by serum testing for detection of virus, viral nucleic acids, or virus-specific IgM and neutralizing antibodies. “During the first week after onset of symptoms, chikungunya can often be diagnosed by performing viral culture or nucleic acid amplification on serum. Virus-specific IgM and neutralizing antibodies normally develop toward the end of the first week of illness. Therefore, to definitively rule out the diagnosis, convalescent-phase samples should be obtained from patients whose acute-phase samples test negative. Testing for chikungunya virus is performed at CDC, several state health department laboratories, and several commercial laboratories” (Staples et al., 2017, 2020).

West Nile Virus (WNV) (Nasci et al., 2013): “WNV infections are most frequently confirmed by detection of anti-WNV immunoglobulin (Ig) M antibodies in serum or cerebrospinal fluid (CSF). The presence of anti-WNV IgM is usually good evidence of recent WNV infection but may indicate infection with another closely related flavivirus (e.g., St. Louis encephalitis). Because anti-WNV IgM can persist in some patients for > 1 year, a positive test result occasionally may reflect past infection unrelated to current disease manifestations. Serum collected within 8 days of illness onset may lack detectable IgM, and the test should be repeated on a convalescent-phase sample. IgG antibody generally is detectable shortly after the appearance of IgM and persists for years. Plaque-reduction neutralization tests (PRNT) can be performed to measure specific virus-neutralizing antibodies. A fourfold or greater rise in neutralizing antibody titer between acute- and convalescent-phase serum specimens collected 2 to 3 weeks apart may be used to confirm recent WNV infection and to discriminate between cross-reacting antibodies from closely related flaviviruses.” NAAT may not be suitable in most cases since the concentrations of WNV RNA are so low by the time a patient begins to show symptoms of infection; however, NAAT may be suitable in immunocompromised individuals who have either delayed or absent antibody development.  

Yellow Fever Virus (YFV) (Gershman & Staples, 2021): Isolation of the virus or NAAT should be performed as early as possible in suspected cases of YFV. “By the time more overt symptoms are recognized, the virus or viral RNA may no longer be undetectable. Therefore, virus isolation and nucleic acid amplification should not be used to rule out a diagnosis of YF … Serologic assays to detect virus-specific IgM and IgG antibodies (sic). Because of cross-reactivity between antibodies raised against other flaviviruses, more specific antibody testing, such as a plaque reduction neutralization test, should be done to confirm the infection (Gershman & Staples, 2021).” Since YFV is a nationally notifiable disease, clinicians should contact their state and/or local health departments according to their respective local, state, and/or federal guidelines. As of April of 2021, “Sanofi Pasteur announced that YF-VAX (yellow fever vaccine) is once again available for purchase in the United States. Providers with a current Yellow Fever Vaccination Stamp issued by their state or territorial health department may now order YF-VAX from the manufacturer” (Gershman & Staples, 2021). 

Dengue (CDC, 2020, 2023c): Diagnosis of dengue can be via isolation of virus, serological tests such as immunoassays, and molecular methods, including RT-PCR. The CDC recommends testing symptomatic individuals that have recently traveled to or lives in areas where dengue is transmitted, as well as symptomatic pregnant women with possible dengue or Zika exposure. The CDC states that dengue virus testing is not recommended for asymptomatic patients and is not recommended for preconception screening. The CDC’s testing algorithm for dengue is as follows:

“Patients with symptoms consistent with dengue can be tested with both molecular and serologic diagnostic tests during the first 7 days of illness. After the first 7 days of illness, test only with serologic diagnostic tests” (CDC, 2020).

Zika / Dengue Virus (CDC): The CDC released updated guidelines associated with Zika and Dengue testing for pregnant individuals. For asymptomatic pregnant individuals “living in or with recent travel to the U.S. and its territories, routine Zika virus testing is not currently recommended . . . recent travel to an area with risk of Zika outside the U.S. and its territories, Zika virus testing is not routinely recommended, but NAAT testing may still be considered up to 12 weeks after travel . . . Zika virus serologic testing is not recommended . . . There is notable cross-reactivity between dengue IgM and Zika IgM antibodies in serologic tests. Antibodies generated by a recent dengue virus infection can cause the Zika IgM to be falsely positive.” 

For symptomatic pregnant individuals who had “recent travel to areas with active dengue transmission and a risk of Zika, specimens should be collected as soon as possible after the onset of symptoms up to 12 weeks after symptom onset. The following diagnostic testing should be performed at the same time: Dengue and Zika virus NAAT testing on a serum specimen, Zika virus NAAT on a urine specimen, and IgM testing for dengue only. Zika virus IgM testing is not recommended for symptomatic pregnant individuals. Zika IgM antibodies can persist for months to years following infection. Therefore, detecting Zika IgM antibodies might not indicate a recent infection. There is also notable cross-reactivity between dengue IgM and Zika IgM antibodies in serologic tests. Antibodies generated by a recent dengue virus infection can cause the Zika IgM to be falsely positive. If the Zika NAAT is positive on a single specimen, the Zika NAAT should be repeated on newly extracted RNA from the same specimen to rule out false-positive NAAT results. If the dengue NAAT is positive, this provides adequate evidence of a dengue infection, and no further testing is indicated. If the IgM antibody test for dengue is positive, this is adequate evidence of a dengue infection, and no further testing is indicated.” 

For symptomatic pregnant individuals who have had sex with someone who lives in or recently traveled to areas with a risk of Zika, specimens should be collected as soon as possible after the onset of symptoms up to 12 weeks after symptom onset. Only Zika NAAT should be performed. If the Zika NAAT is positive on a single specimen, the Zika NAAT should be repeated on newly extracted RNA from the same specimen to rule out false-positive NAAT results.” 

The following should be considered guidance for pregnant individuals who have a fetus with prenatal ultrasound findings consistent with congenital Zika virus infection who live in or traveled to areas with a risk of Zika during her pregnancy. “Zika virus NAAT and IgM testing should be performed on maternal serum and NAAT on maternal urine. If the Zika virus NAATs are negative and the IgM is positive, confirmatory PRNTs should be performed against Zika and dengue. If amniocentesis is being performed as part of clinical care, Zika virus NAAT testing of amniocentesis specimens should also be performed and results interpreted within the context of the limitations of amniotic fluid testing. It is unknown how sensitive or specific RNA NAAT testing of amniotic fluid is for congenital Zika virus infection or what proportion of infants born after infection will have abnormalities. Testing of placental and fetal tissues may also be considered." 

Per the CDC, “Symptomatic non-pregnant patients should refer to testing guidance for dengue. Zika testing is NOT currently recommended for this group based on the current epidemiology of these viruses . . . As per previous guidance, asymptomatic non-pregnant patients should NOT be tested for dengue or Zika viruses. Zika virus testing should NOT be performed as part of preconception screening” (CDC, 2022f). 

The CDC also notes that: “Laboratory testing for congenital Zika virus infection is recommended for infants born to mothers with laboratory evidence of Zika virus infection during pregnancy, and for infants who have abnormal clinical findings suggestive of congenital Zika virus syndrome and a maternal epidemiologic link suggesting possible transmission, regardless of maternal Zika virus test results” (CDC, 2018a).

Infectious Diseases Society of America (IDSA) and the American Society for Microbiology (ASM) 
Laboratory Diagnosis of Tick-borne Infections: The information given below outlines the diagnostic procedures for tick-borne infections and is taken from Table 47 of the 2018 IDSA/ASM guidelines.

The IDSA/ASM does note that most PCR-based assays for babesiosis only detect B. microti even though there are at least three other species of Babesia that can cause the infection. “Real time PCR available from CDC and reference labs… Serology does not distinguish between acute and past infection” (Miller et al., 2018).

Their recommendation for the main diagnostic testing for malaria due to Plasmodium falciparum, P. ovale, P. vivax, P. malariae, and P. knowlesi is “Stat microscopic examination of Giemsa-stained thick and thin blood films (repeat testing every 12 – 24 h for a total of 3 exams before ruling out malaria); rapid antigen detection tests followed by confirmatory blood films within 12 – 24 h.” They make the following special remark: “Antigen tests lack sensitivity with low parasitemia and non-falciparum malaria and do not differentiate all species. PCR from some reference laboratories will detect and differentiate all species. Calculation of percentage parasitemia (using thick or thin blood films) is required for determining patient management and following response to therapy (Miller et al., 2018).” Concerning dengue virus DENV, “Plaque reduction neutralization tests (PRNTs) are considered the reference standard for detection of antibodies to arthropod-borne viruses (arboviruses) and provide improved specificity over commercial serologic assays; however, due to the complexity of testing, PRNT is currently only available at select public health laboratories and the CDC.” They note that false positives for antibodies to DENV may not necessarily indicate DENV infection since it can also be indicative of a prior flavivirus infection, such as West Nile virus or Zika virus. They also state that the “Detection of DENV RNA by NAAT is preferred for acutely ill patients. Recently, detection of the DENV NS1 antigen, which is secreted from infected host cells as early as 1 day after symptom onset and up to 10 days thereafter, has become an acceptable alternative to NAAT for diagnosis of acute DENV infection” (Miller et al., 2018).

For West Nile Virus (WNV), they state: “Laboratory diagnosis of WNV, and most other arboviruses, is typically accomplished by detecting virus-specific IgM- and/or IgG-class antibodies in serum and/or CSF.” Possible false positives can occur if a patient has been vaccinated against yellow fever or if they have had a previous infection of another flavivirus. They do note that WNV RNA detection via NAAT can be performed on either the serum or CSF for immunosuppressed patients.

World Health Organization (WHO)
Interim guidance for laboratory testing of Zika and dengue virus published in July 2022 by WHO includes these updated key considerations, recommendations, and good practices:

• ZIKV and DENV infections need to be differentiated from each other, and from other circulating arboviral and non-arboviral pathogens, using laboratory tests. 
• Laboratory tests performed and interpretation of results must be guided by the interval between symptom onset or exposure, and the collection of specimens. 
• WHO recommends the use of whole blood, serum, or plasma routine diagnostic testing for arboviruses, and urine for ZIKV NAAT testing.
• Molecular assays are the preferred detection method but the period of RNA detectability following infection is limited. 
• Interpretation of serologic test results remains challenging because of cross-reactivity and prolonged detection of virus-specific antibodies; their utility depends on the patient’s current and prior flavivirus exposures. 
• Testing for antibodies to ZIKV and DENV should thus be done with careful consideration of epidemiologic and clinical context. 
• For pregnant women, the diagnosis of ZIKV should always be based on laboratory evidence and testing in these patients should not be limited to a subset of samples, even during outbreaks. 
• For pregnant women, accurate diagnosis is of particular importance; prolonged detection of RNA in blood and urine may facilitate. confirmation of ZIKV infection in these patients 
• ZIKV IgM testing in pregnant women should be used with caution, since a positive test might reflect infection that occurred prior to pregnancy 
• ZIKV testing for asymptomatic pregnant women remains challenging because of unknown optimal timing of specimen collection and risks of false positive and false negative results. 
• Only laboratory tests that have undergone independent, comprehensive assessment of quality, safety and performance should be used for diagnosing arboviral infections. 
• Any testing for the presence of ZIKV, DENV, and other pathogens in the differential diagnosis should be performed in appropriately equipped laboratories by staff trained in the relevant technical and safety procedures (WHO, 2022)

American Society for Microbiology (ASM) 
The ASM updated guidelines in 2022 on laboratory testing for Zika virus. They state, “Diagnostic testing may be warranted for patients who live in or have recently travelled to an endemic region and are critically ill, hospitalized or pregnant, or infants born to Zika virus positive mothers” (ASM, 2022). The ASM endorses CDC guidelines on Zika as well. 

American Academy of Pediatrics 2021-2024 Redbook 
Babesiosis (AAP, 2021a): “Acute, symptomatic cases of babesiosis typically are diagnosed by microscopic identification of Babesia parasites on Giemsa- or Wright-stained blood smears … If the diagnosis of babesiosis is being considered, manual (nonautomated) review of blood smears for parasites should be requested explicitly. If seen, the tetrad (Maltese-cross) form is pathognomonic. B microti and other Babesia species can be difficult to distinguish … examination of blood smears by a reference laboratory should be considered for confirmation of the diagnosis.” They do state that antibody testing can be useful in distinguishing between Babesia and Plasmodium infections whenever blood smear examinations and travel histories are inconclusive or for detecting individuals with very low levels of parasitemia.

Non-Lyme Borrelia Infections (AAP, 2021b): Dark-field microscopy and Wright-, Giemsa-, or acridine orange-stained preparations of blood smears can be used to observe the presence of spirochetes in the initial febrile episode, but their presence is more difficult to determine in future recurrences. Both enzyme immunoassay and Western immunoblot analysis can detect serum antibodies; however, “Antibody tests are not standardized and are affected by antigenic variations among and within Borrelia species and strains.” As of publication, PCR and antibody-based testing were still under development and were not widely available.

Ehrlichia, Anaplasma, and Related Infections (AAP, 2021e): PCR testing should be performed within the first week of illness to diagnose anaplasmosis, ehrlichiosis, and other Anaplasmataceae infections because doxycycline treatment rapidly decreases the sensitivity of PCR. Consequently, negative PCR results do not necessarily indicate a lack of infection. Occasionally, Giemsa- or Wright staining of blood smears can be performed to identify the presence of the morulae of Anaplasma in the first week of illness. Culture testing for isolation is not performed. “Serologic testing may be used to demonstrate a fourfold change in immunoglobulin (Ig) G-specific antibody titer by indirect immunofluorescence antibody (IFA) assay between paired acute and convalescent specimens taken 2 to 4 weeks apart. A single mildly elevated IgG titer may not be diagnostic, particularly in regions with high prevalence. IgM serologic assays are prone to false-positive reactions, and IgM can remain elevated for lengthy periods of time, reducing its diagnostic utility.”

Rocky Mountain Spotted Fever (RMSF) (AAP, 2021h): “The gold standard confirmatory test is indirect immunofluorescence antibody (IFA) to R rickettsii antigen. Both immunoglobulin (Ig) G and IgM antibodies begin to increase around 7 to 10 days after onset of symptoms; IgM is less specific, and IgG is the preferred test. Confirmation requires a fourfold or greater increase in antigen-specific IgG between acute (first 1 – 2 weeks of illness while symptomatic) and convalescent (2 – 4 weeks later) sera.”
Rickettsialpox (AAP, 2021g): Rickettsialpox can be mistaken for other rickettsial infections. Ideally, the use of R. akari-specific antigen is recommended for serologic diagnosis, but it has limited availability. Otherwise, indirect IFA for R. rickettsia, the causative agent of RMSF, since R. akari has extensive cross-reactivity. Again, a demonstration of at least a fourfold increase in antibody titers taken 2-6 weeks apart is indicative of infection.

Chikungunya (AAP, 2021c): “Laboratory diagnosis generally is accompanied by testing serum to detect virus, viral nucleic acid, or virus-specific immunoglobulin (Ig) M and neutralizing antibodies.” RT-PCR can be used to diagnose chikungunya during the first week after onset of symptoms since chikungunya-specific antibodies have not formed at that time. After the first week, serum testing of IgM or a plaque-reduction neutralization test can be performed.

Dengue (AAP, 2021d): “Dengue virus is detectable by RT-PCR or NS1 antigen EIAs from the beginning of the febrile phase until day 7 to 10 after illness onset.” Cross-reactivity occurs between anti-dengue virus IgM and other flaviviruses, including Zika. IgG EIA and hemagglutination testing is not specific for diagnosis of dengue, and IgG antibodies remain elevated for life; consequently, a fourfold increase in IgG between the acute and convalescent phase can confirm recent infection, with “Reference testing is available from the Dengue Branch of the Centers for Disease Control and Prevention.”

Malaria (AAP, 2021f): Microscopic identification of Plasmodium on both thick and thin blood films should be performed. “If initial blood smears test negative for Plasmodium species but malaria remains a possibility, the smear should be repeated every 12 to 24 hours during a 72-hour period … Serologic testing generally is not helpful, except in epidemiologic surveys … Species confirmation and antimalarial drug resistance testing are available free of charge at the Centers for Disease Control and Prevention (CDC) for all cases of malaria diagnosed in the United States.” One FDA-approved RADT is available in the U.S. to hospitals and commercial labs; however, both positive and negative test results must be corroborated by microscopic examination.

West Nile Virus (WNV) (AAP, 2021i): PCR is not recommended for diagnosis of WNV in immunocompetent patients since WNV RNA is usually no longer detectable by the initial onset of symptoms. “Detection of anti-WNV immunoglobulin (Ig) M antibodies in serum or CSF is the most common way to diagnose WNV infection.” Anti-WNV IgM levels can remain elevated for longer than 1 year so a positive test result may be indicative of a prior infection. “Plaque-reduction neutralization tests can be performed to measure virus-specific neutralizing antibodies and to discriminate between cross-reacting antibodies from closely related flaviviruses. A fourfold or greater increase in virus-specific neutralizing antibodies between acute-and convalescent-phase serum specimens collected 2 or 3 weeks apart may be used to confirm recent WNV infection.”

International Encephalitis Consortium (IEC) 
In 2013, the IEC released their Case Definitions, Diagnostic Algorithms, and Priorities in Encephalitis. Concerning arboviruses, they state the following: “For most arboviruses, serologic testing of serum and CSF is preferred to molecular testing, since the peak of viremia typically occurs prior to symptom onset. For example, in patients with West Nile virus (WNV) associated with neuroinvasive disease, CSF PCR is relatively insensitive (57%) compared with detection of WNV IgM in CSF. The cumulative percentage of seropositive patients increases by approximately 10% per day during the first week of illness suggesting the need for repeat testing if the suspicion for disease is strong in those with initially negative results. Notably, arbovirus IgM antibodies may be persistently detectable in the serum and, less commonly, in the CSF, for many months after acute infection, and therefore may not be indicative of a current infection. Therefore, if possible, documentation of acute infection by seroconversion and/or 4-fold or greater rises in titre using paired sera is recommended” (Venkatesan et al., 2013).

Table of Terminology 

References  

1. AAP, A. A. o. P. (2021a). Babesiosis. In D. Kimberlin, M. Brady, M. Jackson, & S. Long (Eds.), Red Book: 2021- 2024 Report of the Committee on Infectious Diseases (pp. 235-237). American Academy of Pediatrics. https://redbook.solutions.aap.org/chapter.aspx?sectionid=189640045&bookid=2205 
2. AAP, A. A. o. P. (2021b). Borrelia Infections Other Than Lyme Disease. In D. Kimberlin, M. Brady, M. Jackson, & S. Long (Eds.), Red Book: 2021- 2021 Report of the Committee on Infectious Diseases (pp. 252-255). American Academy of Pediatrics. https://redbook.solutions.aap.org/chapter.aspx?sectionid=189640055&bookid=2205 
3. AAP, A. A. o. P. (2021c). Chikungunya. In D. Kimberlin, M. Brady, M. Jackson, & S. Long (Eds.), Red Book: 2021-2024 Report of the Committee on Infectious Diseases (pp. 271-272). American Academy of Pediatrics. https://redbook.solutions.aap.org/chapter.aspx?sectionId=189640062&bookId=2205 
4. AAP, A. A. o. P. (2021d). Dengue. In D. Kimberlin, M. Brady, M. Jackson, & S. Long (Eds.), Red Book: 2021-2024 Report of the Committee on Infectious Diseases (pp. 317-319). American Academy of Pediatrics. https://redbook.solutions.aap.org/chapter.aspx?sectionId=189640081&bookId=2205 
5. AAP, A. A. o. P. (2021e). Ehrlichia, Anaplasma, and Related Infections. In D. Kimberlin, M. Brady, M. Jackson, & S. Long (Eds.), Red Book: 2021-2024 Report of the Committee on Infectious Diseases (pp. 323-328). American Academy of Pediatrics. https://redbook.solutions.aap.org/chapter.aspx?sectionid=189640084&bookid=2205 
6. AAP, A. A. o. P. (2021f). Malaria. In D. Kimberlin, M. Brady, M. Jackson, & S. Long (Eds.), Red Book: 2021-2024 Report of the Committee on Infectious Diseases (pp. 527-537). American Academy of Pediatrics. https://redbook.solutions.aap.org/chapter.aspx?sectionId=189640129&bookId=2205 
7. AAP, A. A. o. P. (2021g). Rickettsialpox. In D. Kimberlin, M. Brady, M. Jackson, & S. Long (Eds.), Red Book: 2021-2024 Report of the Committee on Infectious Diseases (pp. 696-697). American Academy of Pediatrics. https://redbook.solutions.aap.org/chapter.aspx?sectionId=189640173&bookId=2205 
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Coding Section  

Procedure and diagnosis codes on Medical Policy documents are included only as a general reference tool for each policy. They may not be all-inclusive. 

This medical policy was developed through consideration of peer-reviewed medical literature generally recognized by the relevant medical community, U.S. FDA approval status, nationally accepted standards of medical practice and accepted standards of medical practice in this community, Blue Cross Blue Shield Association technology assessment program (TEC) and other nonaffiliated technology evaluation centers, reference to federal regulations, other plan medical policies, and accredited national guidelines.

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