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, 2024s). 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., 2024).

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 nucleic acid amplification testing (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 a relapsing fever caused by a Borrelia spp., the following testing is considered MEDICALLY NECESSARY:
   1. For individuals suspected of having hard tick relapsing fever (HTRF) (see Note 2): serologic assays to detect Borrelia antibodies or PCR testing to detect Borrelia miyamotoi.
   2. For individuals suspected of having louse-borne relapsing fever (LBRF) (see Note 3): peripheral blood smear microscopy or PCR testing to detect Borrelia recurrentis.
   3. For individuals suspected of having a soft tick relapsing fever (STRF)/tickborne relapsing fever (TBRF) (see Note 4): dark-field microscopy of a peripheral blood smear, microscopy of a Wright- or Giemsa-stained blood smear, PCR testing to detect Borrelia spp., or serologic assays to detect Borrelia antibodies.
4. For individuals suspected of having a relapsing fever caused by a Borrelia spp., culture testing for Borrelia is considered NOT MEDICALLY NECESSARY.
5. For individuals suspected of having chikungunya (see Note 5), 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.
6. For individuals suspected of having Colorado tick fever (CTF) (see Note 6), the use of PCR testing or IFA for CTF-specific IgM antibodies is considered MEDICALLY NECESSARY.
7. 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 a DENV infection (see Note 7).
   2. For individuals who are symptomatic for Zika virus infection (see Note 8).
8. For individuals suspected of having DENV (see Note 7), the use of IgG ELISA or hemagglutination testing is considered NOT MEDICALLY NECESSARY.
9. For individuals suspected of having ehrlichiosis and/or anaplasmosis (see Note 8), the use of NAAT of whole blood, IFA assay for IgG antibodies, or microscopy for morulae detection is considered MEDICALLY NECESSARY.
10. For individuals suspected of having ehrlichiosis and/or anaplasmosis (see Note 8), the use of an IFA assay for IgM antibodies or standard blood culture is considered NOT MEDICALLY NECESSARY.
11. For individuals suspected of having malaria (see Note 10), 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.
12. To confirm the species of Plasmodium in an individual diagnosed with malaria, PCR testing is considered MEDICALLY NECESSARY.
13. For individuals suspected of having malaria (see Note 10), the use of IFA for Plasmodium antibodies is considered NOT MEDICALLY NECESSARY.
14. For individuals suspected of having a rickettsial disease (see Note 11), the use of an IFA assay for IgG antibodies (two tests occurring a minimum of two weeks apart) is considered MEDICALLY NECESSARY.
15. For individuals suspected of having a rickettsial disease (see Note 11), the use of standard blood culture, NAAT, or IFA assay for IgM antibodies is considered NOT MEDICALLY NECESSARY.
16. For individuals suspected of having West Nile virus (WNV) disease (see Note 12), the use of IFA for WNV-specific IgG or IgM antibodies in either serum or CSF and a confirmatory plaque reduction neutralization test for WNV is considered MEDICALLY NECESSARY.
17. To confirm a WNV infection in individuals who are immunocompromised, nucleic acid detection of WNV is considered MEDICALLY NECESSARY.
18. For immunocompetent individuals suspected of having WNV disease (see Note 12), the use of NAAT for WNV is considered NOT MEDICALLY NECESSARY.
19. For individuals suspected of having a yellow fever virus (YFV) infection (see Note 13), 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.
20. 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 symptoms for symptomatic (see Note 8) pregnant individuals who, during pregnancy, have either lived in or traveled to areas with current or past Zika transmission or who have had sex with someone who either lives in or has recently traveled to areas with current or past Zika virus transmission (see Note 14).
    2. For symptomatic non-pregnant individuals living in or with recent travel to an area with an active CDC Zika Travel Health Notice or an area with current or past Zika virus transmission (see Note 14) when symptoms presented within the last seven days.
21. Zika virus NAAT and Zika virus IgM testing, as well as a confirmatory plaque reduction neutralization test for Zika, is considered MEDICALLY NECESSARY in any of the following situations:
    1. Up to 12 weeks after the onset of symptoms for symptomatic (see Note 8) pregnant individuals who, during pregnancy, have either lived in or traveled to areas with an active CDC Zika Travel Health Notice or who have had sex with someone who either lives in or has recently traveled to areas with an active CDC Zika Travel Health Notice (see Note 14).
    2. For pregnant individuals who have a fetus with prenatal ultrasound findings consistent with congenital Zika virus infection (see Note 15).
    3. For infants born from individuals who, during pregnancy, tested positive for Zika virus.
    4. For infants born with signs and symptoms of congenital Zika syndrome (see Note 15) and who have a birthing parent who had a possible Zika virus exposure during pregnancy.
    5. For symptomatic non-pregnant individuals living in or with recent travel to an area with an active CDC Zika Travel Health Notice or an area with current or past Zika virus transmission (see Note 14) when symptoms presented more than seven days prior to testing.
22. For non-pregnant individuals who have not traveled outside of the United States and its territories and who are symptomatic for Zika virus infection (see Note 8), NAAT and/or IgM testing for Zika detection is considered NOT MEDICALLY NECESSARY.
23. 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, 2024j).

Note 2: Typical signs and symptoms of HTRF (caused by Borrelia miyamotoi) can include chills or shakes, fatigue, nausea or vomiting, headache, and muscle and joint aches (CDC, 2024a).

Note 3: Typical signs and symptoms of LBRF (caused by Borrelia recurrentis) can include fever, headache, chills or shakes, muscle and joint aches, and nausea. Though the clinical symptoms of LBRF are similar to STRF, LBRF is usually associated with fewer relapses (CDC, 2024b)

Note 4: Typical signs and symptoms of STRF/TBRF (caused by Borrelia hermsii, B. turicatae, and other Borrelia bacteria) can include fever, headache, muscle aches, chills, dizziness, joint pain, nausea and vomiting, appetite loss, and rarely, facial paralysis eye pain or redness, or vision changes (CDC, 2024c).

Note 5: 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., 2024).

Note 6: Typical signs and symptoms of CTF can include fever, chills, headache, myalgia, malaise, sore throat, vomiting, abdominal pain, and maculopapular or petechial rash (CDC, 2024e).

Note 7: Typical signs and symptoms of dengue include fever, headache, retro-orbital eye pain, myalgia, arthralgia, macular or maculopapular rash, petechiae, ecchymosis, purpura, epistaxis, gingival bleeding, hematuria, leukopenia, thrombocytopenia, hyponatremia, elevated AST and ALT, and nausea and/or vomiting (CDC, 2024f, 2024r).

Note 8: Typical signs and symptoms of Zika virus infection can include fever, rash, headache, joint pain, conjunctivitis (red eyes), and muscle pain (CDC, 2024t).

Note 9: 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 10: 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 (Tan & Abanyie, 2024).

Note 11: 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 12: 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 13: 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, 2024).

Note 14: The CDC provides information on the geographic risk classifications of Zika (https://www.cdc.gov/zika/geo/index.html), as well as providing travel health notices for pathogens of concern (https://wwwnc.cdc.gov/travel/notices). 

Note 15: 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, 2024n).

Table of Terminology

Term	Definition
AAP	American Academy of Pediatrics
ASM	American Society for Microbiology
CDC	Centers for Disease Control and Prevention
CMS	Centers for Medicare & Medicaid Services
CSF	Cerebrospinal fluid
CTF/CTFV	Colorado tick fever /virus
CV	Coefficient of variation
DENV	Dengue virus
DENV NS1	Dengue virus nonstructural protein 1
DHF	Dengue hemorrhagic fever
DNA	Deoxyribonucleic acid
EDTA	Ethylenediaminetetraacetic acid
EIA	Enzyme immunoassay
ELISA	Enzyme-linked immunosorbent assays
EM	Erythema migrans
ESR	Erythrocyte sedimentation rate
FDA	Food and Drug Administration
FFPE	Formalin-fixed, paraffin-embedded
FISH	Fluorescent in situ hybridization
GlpQ	Glycerophosphodiester phosphodiesterase gene
HAI	Hemagglutination inhibition test
HTRF	Hard tick relapsing fever
IDSA	Infectious Diseases Society of America
IEC	International Encephalitis Consortium
IFA	Indirect immunofluorescence antibody
IFAs	Immunofluorescence assays
IgG	Immunoglobulin G
IgM	Immunoglobulin M
IHC	Immunohistochemistry
IMCA	Immunochemiluminometric assay
LBRF	Louse-borne relapsing fever
LDTs	Laboratory developed tests
MAbs	Monoclonal antibodies
MAC-ELISA	IgM antibody capture enzyme-linked immunosorbent assay
MIA	Microsphere-based immunoassay
MIF	Microimmunofluorescent
NAAT	Nucleic acid amplification testing
NDPH	New daily persistent headache
NNDSS	National Notifiable Disease Surveillance System
PCR	Polymerase chain reaction
PRNT	Plaque reduction neutralization test
PRNTs	Plaque reduction neutralization tests
PT	Prothrombin time
PTT	Partial thromboplastin time
qPCR	Quantitative polymerase chain reaction
RDT	Rapid diagnostic testing
RMSF	Rocky Mountain spotted fever
RNA	Ribonucleic acid
RT-PCR	Real-time polymerase chain reaction
SFG	Spotted fever group
STRF	Soft tick relapsing fever
TBRF	Tickborne relapsing fever
WHO	World Health Organization
WNV	West Nile virus
YFV	Yellow fever virus

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, 2022b).

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 tickborne bacteria and protozoa more than doubled from 2004 to 2016. “In the United States, 16 vector-borne 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 tickborne disease increase occurs in the continental U.S. and has experienced a gradual, steady rate increase with Lyme disease comprising 82% of all tickborne 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.

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, 2024q). RMSF is caused by Rickettsia rickettsii, 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, 2024q). 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 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 (McClain, 2024a). 

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 (McClain, 2024c). 

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 (McClain, 2024b). 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 (McClain, 2024b).

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, 2024; Miller et al., 2024). 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 three 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, 2024p). One exception is using antibodies to the GlpQ protein characteristic of these Borrelia species but not to B. burgdorferi (Lyme disease) (Barbour, 2024).

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: one to four 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 tickborne 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, 2024). 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, 2022). 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” Clinic (2024).

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 (Petersen, 2022).

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, 2024f). “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., 2022). 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., 2022). 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, 2024t). The most common symptoms of Zika are fever, rash, joint pain, and conjunctivitis (CDC, 2024t). The illness is usually mild with symptoms beginning two to seven 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, 2024t; LeBeaud, 2023). 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, 2023).

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 (Petersen, 2021).

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 four of ten 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, 2024).

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, 2022).

Types of Testing

Test	Description	Rationale
Culture	Culture growth depends on the pathogen being studied. If the pathogen is an obligate intracellular organism, then it must be isolated using more sophisticated cell culture techniques. In many circumstances, culture is used for research and/or epidemiology rather than as a diagnostic tool (Biggs et al., 2016; Miller et al., 2024).	At times, culture testing is not as sensitive as either NAAT or serologic testing and can be time-intensive when treatment should not be delayed. Depending on the organism, this may require high biosafety level laboratory for culture growth (Biggs et al., 2016).
Indirect immunofluorescene antibody (IFA) assays	IFA is a serologic assay that can be used to test for the presence of antibodies, such as IgG and IgM, reactive against the pathogen (Biggs et al., 2016).	Depending on the pathogen, IFA can be a useful tool. At times, though, it can cross-react with either a prior vaccination or infection (Wilder-Smith, 2024). An acute infection can often be determined by performing IFA in both the acute phase and convalescent phase where at least a fourfold increase in antibodies is indicative of an acute infection (Biggs et al., 2016).
Darkfield microscopy	Darkfield microscopy can be used to detect the presence of microorganisms, such as motile spirochetes (Miller et al., 2024).	This technique is not widely available, and transport of sample must be done immediately if testing of motile specimen is desired (Miller et al., 2024).
Blood-smear microscopy	Blood-smear microscopy can be either thick or thin and is typically performed on a sample stained with an eosin-azure-type dye, such as Giemsa, to look at intracellular structures or morphological features (Biggs et al., 2016).	This technique should be performed by an experienced microscopist since it can be inconsistent. As compared to other techniques, this technique is relatively inexpensive (Biggs et al., 2016).
Nucleic acid amplification testing (NAAT)	NAATs can include polymerase chain reaction (PCR), real-time PCR (RT-PCR), or other enzyme-dependent amplification testing for the presence of nucleic acids (DNA or RNA).	NAATs can be specific and sensitive; however, they may not be available at all laboratories and/or can be costly. Some NAATs are available as rapid diagnostic tools. NAATs have been used on serum, whole blood, tissue, CSF, and even formalin-fixed, paraffin-embedded biopsies from autopsy tissues. The sensitivity of the technique can vary depending on the sample; for example, whole blood PCR for R. rickettsii is less sensitive than a similar sample test for E. chaffeensis (Biggs et al., 2016).

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 et al. (2001) 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 the 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 films 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
Kato et al. (2013) 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 (one hour versus one to two 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 (five percent). 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 two 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 Morbidity and Mortality Weekly Report. 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 two to four 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).

In 2024, the CDC published updated guidelines pertaining to rickettsial infections, which provide similar guidelines to those published in 2016. “The standard serologic test for diagnosis of RMSF is the indirect fluorescent antibody (IFA) test for immunoglobulin G (IgG) using R. rickettsii antigen. IgG IFA assays should be performed on paired acute and convalescent serum samples collected 2–10 weeks apart to demonstrate evidence of a fourfold seroconversion. ... Single or inappropriately timed serologic tests, in relation to clinical illness, can lead to misinterpretation of results” (CDC, 2024d). They also provide statements on nucleic acid testing and IHC/culture testing for rickettsial infections: “PCR amplification is performed on DNA extracted from whole blood serum, or plasma. R. rickettsii infect the endothelial cells that line blood vessels and may not circulate in large numbers in the blood until the disease has progressed to a severe phase of infection. Although a positive PCR result is helpful, a negative result does not rule out the diagnosis, and treatment should not be withheld due to a negative result. PCR might also be used to amplify DNA from a skin biopsy of a rash lesion, or in post-mortem tissue specimens. . . Culture and IHC assays can also be performed on skin biopsies of a rash lesion, or post-mortem tissue specimens. Culture isolation and IHC assays of R. rickettsii are only available at specialized laboratories; routine hospital blood cultures cannot detect the organism” (CDC, 2024d).

Soft tick relapsing fever (STRF) /Tickborne relapsing fever (TBRF) (CDC, 2024c, 2024i): In the U.S., STRF/TBRF can be caused by Borrelia hermsii, B. turicatae, and other Borrelia bacteria via the bite of soft-bodied Ornithodoros genus ticks. STRF often presents with a relapsing nature, with symptoms appearing 4 – 21 days after exposure, with intermittent fevers lasting for three days and remitting for seven days before relapse. Moreover, “Spirochetes may be present in high concentrations in the blood of febrile patients (> 106 spirochetes/ml). Spirochetes are most readily detected by microscopy in symptomatic, untreated patients early in the course of infection. Direct visualization by microscopy using dark field or stained peripheral blood smears is generally adequate to confirm the diagnosis. … PCR is more sensitive than microscopy and may also be used during asymptomatic periods or soon after treatment initiation. The preferred specimen type for PCR testing is whole blood. … Serologic testing is available from some labs to diagnose STRF. Serologic assay results are most sensitive when specimens are collected at least 14 days after symptom onset. … Patients with relapsing fevers might have false positive serologic tests for Lyme disease” (CDC, 2024c).

The CDC acknowledges that some PCR and serologic tests may cross-react with other Borrelia species; thus, “clinical and epidemiologic features, such as travel and exposure history, are important to guide interpretation of test results. Consider a diagnosis of STRF for patients with positive Lyme disease or [hard tickborne relapsing fever] serology who have not been in areas endemic for these diseases.” Additionally, patients may exhibit other general laboratory findings, such as “thrombocytopenia, increased white blood cell count, mildly increased serum bilirubin level, elevated erythrocyte sedimentation rate (ESR), and slightly prolonged prothrombin time (PT) and partial thromboplastin time (PTT) (CDC, 2024i).

Hard tick relapsing fever (HTRF) (CDC, 2024a, 2024g): In the U.S., HTRF is used to differentiate between infections caused by hard-bodied ticks and soft-bodied ticks (see STRF above). HTRF is caused by the Borrelia miyamotoi bacteria and is transmitted through the bites of infected blacklegged ticks (Ixodes scapularis) and western blacklegged ticks (Ixodes pacificus). Unlike STRF, it causes a single episode of fever more commonly, with 10% of cases having a relapsing fever. Symptoms appear about two weeks after a tick bite but can occur within three to six days after exposure. Diagnosis is often made by PCR using whole blood, but several PCR and serologic methods cannot distinguish between HTRF and STRF. The CDC also adds “Serologic testing is available from some labs for diagnoses of HTRF. Serologic assay results are most sensitive when specimens are collected at least 14 days after symptom onset. Serum taken early during infection may yield negative results.” Similar emphasis is placed on considering clinical and epidemiological features when interpreting results, as HTRF patients may also test positive for other Borrelia species, such as Lyme disease (CDC, 2024a, 2024g).

Louse-borne relapsing fever (LBRF) (CDC, 2024b, 2024h): In the U.S., LBRF is caused by Borrelia recurrentis bacteria and transmitted by the human body louse, and rarely, head louse. It also occurs endemically in regions of Africa and in overcrowded conditions. Clinically, LBRF presents similarly to STRF but with fewer relapses. Diagnosis is made with “direct visualization of spirochetes in a peripheral blood smear in symptomatic, untreated patients early in the course of infection,” as “people with LBRF experience high levels of spirochetemia during febrile episodes.” Alternatives for diagnosis also include PCR, but the same precautions hold for LBRF as for HTRF and STRF when interpreting results (CDC, 2024b, 2024h). 

Colorado Tick Fever (CTF) (CDC, 2024e): As of 2023, CTF was reportable in Arizona, Colorado, Idaho, Montana, New Mexico, Oregon, South Dakota, Utah, Washington, 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, 2024e). 

Babesiosis (CDC, 2024j): Babesiosis is caused most commonly by Babesia microti, which is usually transmitted by white-footed mice and other small mammals. 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. When considering a babesiosis diagnosis, healthcare providers should explicitly request a manual (non-automated) review of the peripheral blood smear. In symptomatic patients with acute infection, it is typical to detect Babesia parasites through light-microscopic examination of blood smears, though multiple smears may need to be examined. Distinguishing between Babesia and Plasmodium (especially P. falciparum) parasites and artifacts like stain or platelet debris can be challenging. 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, 2024j). 

Malaria (Tan & Abanyie, 2024): 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. Tests should be performed immediately when ordered by a health care provider, and microscopy results should be available as soon as possible, ≤ 24 hours of the patient’s presentation. 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” (Tan & Abanyie, 2024). 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. Furthermore, in some areas, mutations are increasingly being observed in malaria parasites, resulting in an absence of the malaria antigen usually detected by many RDTs, including the only RDT used in the United States. The absence of this parasite antigen in peripheral blood can lead to false-negative RDT test results. 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” (Tan & Abanyie, 2024). Regarding PCR, the CDC states that “These tests are more sensitive than routine microscopy, but 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” (Tan & Abanyie, 2024).

While diagnosis from microscopic examination remains the gold standard for laboratory confirmation of malaria, the CDC does acknowledge that antigen detection with a rapid diagnostic test and molecular diagnosis by PCR may be useful in certain situations: “In the international setting, various test kits are available to detect antigens derived from malaria parasites. Such immunologic ("immunochromatographic") tests most often use a dipstick or cassette format and provide results in 2-15 minutes. These "Rapid Diagnostic Tests" (RDTs) offer a useful alternative to microscopy in situations where reliable microscopic diagnosis is not available. Malaria RDTs are currently used in some clinical settings and programs. On June 13, 2007, the U.S. Food and Drug Administration (FDA) approved the first RDT for use in the United States. This RDT is approved for use by clinical laboratories, not by individual clinicians or by patients themselves. It is recommended that all RDTs are followed-up with microscopy to confirm the results and if positive, to confirm the species and quantify the proportion of red blood cells that are infected. The use of this RDT may decrease the amount of time that it takes to determine whether a patient is infected with malaria. ... Parasite nucleic acids are detected using polymerase chain reaction (PCR). Although this technique may be more sensitive than blood smear microscopy, it is of limited utility for the diagnosis of acutely ill patients in the standard healthcare setting. PCR results are often not available quickly enough to be of value in establishing the diagnosis of malaria infection. PCR is most useful for confirming the species of malarial parasite after the diagnosis has been established by either smear microscopy or RDT” (CDC, 2024k). 

Chikungunya (Staples et al., 2024): In the CDC Yellow Book, concerning the Chikungunya virus, they recommend that “the differential diagnosis of chikungunya virus infection depends on clinical features (signs and symptoms) as well as where the person was suspected of being infected. Consider other diseases in the differential diagnosis, including adenovirus, other alphaviruses (Barmah Forest, Mayaro, O’nyong-nyong, Ross River, and Sindbis), dengue, enterovirus, leptospirosis, malaria, measles, parvovirus, rubella, group A Streptococcus, typhus, Zika, and postinfectious arthritis and rheumatologic conditions. Laboratory diagnosis is done by serum testing for detection of virus, viral nucleic acid, or virus-specific IgM and neutralizing antibodies. Because the virus develops high levels of viremia during the first week after symptom onset, chikungunya can often be diagnosed by performing viral culture or nucleic acid amplification on serum. Virus-specific IgM antibodies normally develop toward the end of the first week of illness but can remain detectable for months to years after infection. Rarely, serum IgM antibody testing can yield false-positive results due to cross-reacting antibodies against related alphaviruses (e.g., Mayaro virus, O’nyong-nyong virus). …Testing for chikungunya virus is performed at several state health department laboratories, and commercial laboratories” (Staples et al., 2024).

West Nile Virus (WNV) (CDC, 2024o): “The front-line screening assay for laboratory diagnosis of human WNV infection is the IgM assay. Currently, the FDA has cleared three commercially available test kits from different manufacturers, for detection of WNV IgM antibodies…In addition, the CDC-defined IgM and IgG EIA [i.e., ELISA or microsphere-based immunoassay (MIA)] can be used. …The CDC MIA can differentiate WNV from St. Louis encephalitis. … Because the IgM and IgG antibody tests can cross-react between flaviviruses (e.g., [St. Louis encephalitis], dengue, yellow fever, WNV, Powassan), they should be viewed as screening tests only. For a case to be considered confirmed, serum samples that are antibody-positive on initial screening should be evaluated by a more specific test; currently the plaque reduction neutralization test (PRNT) is the recommended test for differentiating between flavivirus infections. … Specimens submitted for WNV testing should also be tested against other arboviruses known to be active or be present in the area or in the region where the patient traveled.” 

There are also virus detection assays that can be utilized to detect viable WNV, WNV antigen or WNV RNA in human samples, but they vary in sensitivity, specificity, and time required to conduct the test. However, the CDC warns that “viremia is almost always absent by the time a patient presents with neuroinvasive illness and thus viral isolation is generally not recommended as part of a testing algorithm in immune competent patients. … Confirmation of virus isolate identity can be accomplished by indirect immunofluorescence assay (IFA) using virus-specific monoclonal antibodies (MAbs) or nucleic acid detection (e.g. RT-PCR, real-time RT-PCR or sequencing). … Virus isolation or RT-PCR on serum may be helpful in confirming WNV infection in immunocompromised patients when antibody development is delayed or absent” (CDC, 2024o).

Yellow Fever Virus (YFV) (Gershman & Staples, 2024): 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 detectable; thus, virus isolation and nucleic acid amplification should not be used to rule out a diagnosis of YF. Serologic assays can be used to detect virus-specific IgM and IgG antibodies. Because of the possibility of cross-reactivity between antibodies against other flaviviruses, however, more specific antibody testing (e.g., a plaque reduction neutralization test) should be performed to confirm the infection” (Gershman & Staples, 2024). Since YFV is a nationally notifiable disease, clinicians should contact their state and/or local health departments or call the CDC Arboviral Diseases Branch according to their respective local, state, and/or federal guidelines. As of May 2023, “Only one YF vaccine (YF-VAX, Sanofi Pasteur) is licensed for use in the United States. Periodically in the United States, shortages of YF-VAX have occurred due to production issues, including one that lasted from late 2015 until early 2021. To address this most recent shortage, Sanofi Pasteur collaborated with the CDC and the U.S. Food and Drug Administration (FDA) to import and distribute Stamaril (a YF vaccine comparable to YF-VAX, manufactured at the company’s facility in France) under an expanded-access investigational new drug protocol” (Gershman & Staples, 2024). 

Dengue (CDC, 2024m): Diagnosis of dengue can be diagnosed differently based on the phase: the acute phase (0-7 days after symptom onset) and the convalescent phase (> 7 days after symptom onset). In the acute phase, the CDC recommends diagnosis using one of two testing combinations: “a nucleic acid amplification test (NAAT) (e.g., RT-PCR) and an IgM antibody test OR an NS1 antigen test and an IgM detection test,” but a serum sample is preferred in this stage. However, “a negative result from a RT-PCR or NS1 test does not rule out infection.” Furthermore, the CDC recommends that “when the acute (0 – 7 days) sample is negative in the recommended test combinations or is not available, a convalescent serum sample can be collected and tested.” For the convalescent sample, “IgM ELISA is recommended as the primary test after day 8 of symptom onset;” the CDC warns that after day 7 of illness, NAAT or NS1 antigen tests may not be as sensitive for disease detection. 

The CDC does not recommend serologic testing by IgG for “diagnosis of acute dengue in patients, as these tests may detect antibodies from dengue infections or other flavivirus infections that occurred in the past.” 

With regards to specific circumstances, “for people living in or traveling to an area with concurrently circulating flaviviruses, clinicians will need to order plaque reduction neutralization test (PRNT) to rule out dengue on IgM-positive specimens,” but PRNT does not always give a conclusive diagnostic result, “particularly in patients that have previously been exposed to more than one flavivirus.” Additionally, “if the patient is pregnant and symptomatic and lives in or has traveled to an area with risk of Zika, test for Zika using molecular tests in addition to dengue” (CDC, 2024m).

Zika Virus (CDC, 2024l): The CDC released updated guidelines associated with Zika testing for pregnant individuals. The recommendations for asymptomatic pregnant patients are shown below:

Lived in or traveled to the United States and its territories during pregnancy	Since no confirmed cases of Zika virus have been detected in the United States and its territories since 2018, routine Zika testing is not recommended.
Traveled to an area with an active CDC Zika Travel Health Notice during pregnancy	NAAT testing may be considered up to 12 weeks after travel
Traveled to an area with current or past Zika virus transmission outside the U.S. and its territories during pregnancy	Routine testing is not recommended. If the decision is made to test, NAAT testing can be done up to 12 weeks after travel.

Recommendations for symptomatic pregnant patients are shown below:

Lived in or traveled to an area with an active CDC Zika Travel Health Notice during pregnancy OR had sex during pregnancy with someone living in or with recent travel to an area with an active CDC Zika Travel Health Notice	Specimens should be collected as soon as possible after onset of symptoms up to 12 weeks after symptom onset. Perform dengue and Zika virus NAAT and IgM testing on a serum specimen and Zika virus NAAT on a urine specimen. If Zika NAAT is positive and the Zika IgM is negative, repeat NAAT test on newly extracted RNA from same specimen to rule out false-positive results. If both dengue and Zika virus NAATs are negative but either IgM antibody test is positive, confirmatory PRNTs should be performed against dengue, Zika, and other flaviviruses endemic to the region where exposure occurred.
Lived in or traveled to an area with current or past Zika virus transmission during pregnancy	Specimens should be collected as soon as possible after onset of symptoms up to 12 weeks after symptom onset. Perform dengue and Zika virus NAAT testing on a serum specimen and Zika virus NAAT on a urine specimen. If Zika NAAT is positive, repeat test on newly extracted RNA from same specimen to rule out false-positive results. Perform IgM testing for dengue only. If dengue NAAT or IgM test is positive, this provides adequate evidence of dengue infection, and no further testing is indicated.
Had sex during pregnancy with someone living in or with recent travel to an area with current or past Zika virus transmission	Specimens should be collected as soon as possible after onset of symptoms up to 12 weeks after symptom onset. Only Zika NAAT should be performed. If Zika NAAT is positive, repeat test on newly extracted RNA from same specimen to rule out false-positive results.

For pregnant patients having a fetus with prenatal ultrasound findings consistent with congenital Zika virus infection, the recommendations are below:

Lived in or traveled during pregnancy to areas with an active CDC Zika Travel Health Notice or current or past Zika virus transmission OR had sex during pregnancy with someone living in or with recent travel to areas with an active CDC Zika Travel Health Notice or current or past Zika virus transmission

Zika virus NAAT and IgM testing should be performed on pregnant person's serum and NAAT on pregnant person's 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. Testing of placental and fetal tissues may also be considered.

For symptomatic non-pregnant patients, the recommendations are listed below:

Living in or with recent travel to the United States and its territories

Since no confirmed cases of Zika virus disease have been detected in the United States and its territories since 2018, routine Zika virus testing is not recommended.

Living in or with recent travel to an area with an active CDC Zika Travel Health Notice OR to an area with current or past Zika virus transmission outside the U.S. and its territories

Dengue and Zika virus NAATs should be performed on serum collected ≤7 days after symptom onset. A positive NAAT result typically provides evidence of acute infection. Perform dengue and Zika virus IgM antibody testing on NAAT-negative serum specimens and serum collected >7 days after onset of symptoms. If either dengue or Zika virus IgM antibody testing is positive, and definitive diagnosis is needed for clinical or epidemiologic purposes, confirmatory PRNTs should be performed against dengue, Zika, and other flaviviruses endemic to the region where exposure occurred.

For infants with possible congenital Zika virus infection via gestational parents with possible Zika virus exposure during pregnancy, the CDC recommends to: 

• “Collect specimens as soon as possible after birth.
• Zika virus NAAT and IgM testing should be performed on infant serum and NAAT on infant urine.
• If cerebrospinal fluid (CSF) is obtained for other purposes, NAAT and IgM antibody testing should be performed on CSF.
• If the infant’s serum is IgM non-negative and NAAT negative, but PRNT was not performed on the gestational parent’s serum, PRNT for Zika and dengue viruses should be performed on the infant serum.
• Perform PRNT on a sample collected from an infant aged 18 months or older whose initial sample collected at birth was IgM non-negative and neutralizing antibodies were detected by PRNT in either the infant’s or gestational parent’s sample.”

For asymptomatic non-pregnant patients, “testing for dengue or Zika viruses is not recommended for this group” (CDC, 2024l). 

Infectious Diseases Society of America (IDSA) and the American Society for Microbiology (American Society of Microbiology) 
Laboratory Diagnosis of Tickborne Infections: The information given below outlines the diagnostic procedures for tickborne infections and is taken from Table 50 of the 2024 IDSA/ASM guidelines.