January–June 2017

The Role of Birds in Arboviral Disease Surveillance

in Harris County and the City of Houston, Texas . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 1

Lauren Wilkerson, MS; Martin Reyna Nava, MS; Cheryl Battle-Freeman; et al

Laboratory and Semi-field Evaluations of Two (Transfluthrin) Spatial

Repellent Devices Against Aedes aegypti (L.) (Diptera: Culicidae) . . . . . . . . . . . . . . . . . . . . . . . . 13

MAJ Lee P. McPhatter, USA; Paula D. Mischler, PhD; Meiling Z. Webb, PhD; et al

Georgia’s Collaborative Approach to Expanding Mosquito Surveillance

in Response to Zika Virus: A Case Study . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 23

R. Christopher Rustin, DrPH, MT, REHS; Deonte Martin; Varadan Sevilimedu, MPH; et al

A Location-specific Spreadsheet for Estimating Zika Risk and Timing

for Zika Vector Surveillance, Using US Military Facilities as an Example . . . . . . . . . . . . . . . . . . . 34

Desmond H. Foley, PhD; David B. Pecor

Biosurveillance and Morphological Variations of Larvae and Pupae of Common Malaria

Vectors, Anopheles (Anopheles) Hyrcanus Group Species in the Republic of Korea . . . . . . . . . . 47

Leopoldo M. Rueda, PhD; Heung-Chul Kim, PhD; Sung-Tae Chong, MS; et al

A Case of Chagas Cardiomyopathy Following Infection in South Central Texas . . . . . . . . . . . . . 55

Maj Bryant J. Webber, USAF; Lt Col Edward J. Wozniak, TXSG; CPT David Chang, USA; et al

Direct Detection of Leishmania from Clinical Samples . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 60

John N. Waitumbi, PhD; MAJ Joshua Bast, USA; Nancy Nyakoe, MS; et al

New Records, Distribution, and Updated Checklists of Old World Phlebotomine

Sand Flies, With Emphasis on Africa, Southwest Asia, and Central Asia . . . . . . . . . . . . . . . . . . . 65

Leopoldo M. Rueda, PhD; James E. Pecor; Matthew Wolkoff; et al

Operational Mosquito and Vector-Borne Diseases Surveillance

at Incirlik Air Base, Turkey . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 86

Will K. Reeves, PhD; Myrna M. Miller, DVM, PhD; Orhan Bayik, PHS; Maj Leah Chapman, USAF

Vector-Borne Diseases of Public Health Importance for Personnel

on Military Installations in the United States . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 90

Melissa N. Garcia, PhD, MPH; Thomas L. Cropper, DVM, MPVM, ACVPM; et al

Blast-Associated Traumatic Brain Injury in the Military as a Potential

Trigger for Dementia and Chronic Traumatic Encephalopathy . . . . . . . . . . . . . . . . . . . . . . . . . 102

Jamal Hasoon, MD

Challenges of Practicing Environmental Epidemiology in the US Military . . . . . . . . . . . . . . . . 106

Coleen Baird, MD, MPH; Jessica Sharkey, MPH; Joel Gaydos, MD, MPH

Indirect Military Occupational Lead Exposure to Children at Home: A Case Report . . . . . . . 112

LTC(P) Paul O. Kwon, USA

Spurgeon Neel 2016 Annual Writing Award Winner

Enduring for the Patients’ Sake: The Emotional Experiences and Endurance of American

Ambulance Drivers in World War I . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . . 115

Benjamin Cameron Schaffer

JOURNAL

THE UNITED STATES ARMY

MEDICAL DEPARTMENT

Vector-Borne Diseases and Challenges to Public Health
JOURNAL

A Professional Publication

of the AMEDD Community THE UNITED STATES ARMY

MEDICAL DEPARTMENT

Online issues of the AMEDD Journal are available at http://www.cs.amedd.army.mil/amedd_journal.aspx

January–June 2017 US Army Medical Department Center & School PB 8-17-1/2/3/4/5/6

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January – June 2017 1

The Avian Section of Harris County Public Health 

(HCPH) Mosquito and Vector Control Division (MVCD) 

is responsible for monitoring avian populations in Harris County and the City of Houston, Texas, for West Nile 

virus (WNV), St. Louis encephalitis (SLE), and other 

mosquito-borne diseases. Since birds are the main hosts 

for many of these viruses, they are included in mosquito 

surveillance activities to provide a complete picture of 

the enzootic cycle rather than simply monitoring mosquitoes alone. While infected birds are only viremic for 

a few days,1

 surviving birds produce detectible levels of 

virus specific antibodies for a much longer period, up to 

several years.2,3 This persistence allows for a more longterm “big picture” surveillance effort that can identify 

the presence of a virus in a new area even if no virus is 

currently circulating.

In 1999, the first sign that WNV had reached the United 

States was due to large numbers of dead birds, primarily crows, in New York City.4

 Subsequently, wildlife 

and disease ecologists noted continued large die-offs 

of crows and Blue Jays as the virus spread across the 

continent.5

 Harris County Public Health MVCD created 

a Dead Bird Hotline for residents to report dead birds. 

The first detection of WNV in the Houston area was 

from a dead Blue Jay in 2002.

Since WNV has been established in Harris County, 

avian monitoring helps explain the complex ecological 

interactions responsible for outbreaks. For humans to be 

infected, the virus must first amplify in the mosquitoavian transmission cycle. Avian antibodies, like human 

ones, are important to immune responses and make surviving, seropositive birds essentially resistant to reinfection.6

 In standard epidemiological susceptible-infectedremoved (SIR) models, large numbers of immune individuals reduce the viral transmission rate below a sustainable level, bringing outbreaks to an end.7

 Although 

more research is needed, there is evidence that some 

arboviruses, such as WNV and SLE, follow multiyear 

SIR cycles. A 40-year history of SLE seroprevelances 

in Harris County demonstrated a consistent cyclical 

pattern while a multiyear field study in Chicago found 

that the best predictor of WNV activity in a given year 

was the proportion of seropositive birds at the end of the 

previous year.2

 Similarly, in Los Angeles, a preseason 

seropositive rate of less than 10% was associated with 

WNV outbreaks.8

 This study provides the results from 

the 2015 MVCD avian surveillance activities.

Collection Methods

Live bird trapping was conducted daily throughout the 

year, except when prevented by inclement weather. Trapping locations were located in Houston or Harris County 

parks and varied on a rotational basis. Sites that were 

chosen for live bird trapping provided the most uniform 

coverage available, ie, included historic and current 

WNV and SLE “hot spots” and provided a high degree 

of trapping success. In 2015, 27 sites were trapped, with 

20 monitored biweekly from June to December when 

The Role of Birds in Arboviral Disease

 Surveillance in Harris County and the

 City of Houston, Texas

Lauren Wilkerson, MS Hilda Guzman

Martin Reyna Nava, MS Robert Tesh, MD, PhD

Cheryl Battle-Freeman Mustapha Debboun, PhD, BCE

Amelia Travassos da Rosa

Abstract

Avian arboviral surveillance is an integral part of any disease-based integrated mosquito control program. The 

Harris County Public Health Mosquito and Vector Control Division has performed arboviral surveillance in 

the wild birds of Harris County and the City of Houston since 1965. Blood samples from live trapped birds were 

tested for arboviral antibodies to West Nile virus (WNV), St. Louis encephalitis, Eastern equine encephalitis, 

and Western equine encephalitis. A dead bird surveillance program was created in 2002 with the arrival of 

WNV in Harris County. Since implementation, the program has detected considerable variability in viral activity with annual WNV seroprevelance rates ranging from 2.9% to 17.7%, while the percentage of positive dead 

birds has ranged from 0.3% to 57.2%. In 2015, 1,345 live birds were sampled and 253 dead birds were tested, 

with WNV incidence rates of 16.5% and 5.9%, respectively.
2 http://www.cs.amedd.army.mil/amedd_journal.aspx

seasonal help was available, and on a monthly rotation 

during the rest of the year (Figure 1).

During bird trapping, an average of 3 to 5 Japanese mist 

nets were set up before sunrise and maintained until 

9:30 am. The nets were 2.6 m in height and 6 or 12 m in 

length, and attached to poles that were hammered into 

the ground with mallets. Nets were positioned in areas 

that were expected to attract birds or serve as flyways 

and baited with commercial bird seed at least one day 

prior to trapping. Specific characteristics indicating favorable areas included open spaces interspersed with 

layered vegetation, backyard bird feeders, open trash 

cans, and picnic tables. Nets were checked at least every 15 minutes with optimum trapping occurring when 

all nets were observed from one central location. After trapped birds were removed from the mist nets, they 

were placed in a holding bag, transported to a mobile 

laboratory station and processed on-site.

During processing, we recorded the bird’s species, sex 

(if possible), age, and whether or not it was previously 

banded, as required by the Bird Banding Laboratory. 

We used the National Geographic Society Field Guide 

to Birds of North America9

 along with The Sibley Field 

Guide to Birds of Eastern North America10 for species 

identification, and Pyle’s Identification Guide to North 

American Birds11 for ageing, sexing, and determining 

the correct band size. The age of a bird is of particular 

interest for WNV and SLE surveillance because positive hatch year birds indicate exposures that occurred 

within the current season. After we collected all pertinent information, we drew blood from the bird’s jugular 

vein or brachial vein if it was a dove, because the jugular 

vein is frequently obscured by engorged skin in doves. 

We collected either 0.1 cc of blood for smaller (sparrowsized) birds or 0.2 cc for larger birds (ie, Blue Jays) and 

added it to a corresponding amount of 0.4 cc or 0.8 cc 

diluent of 0.9% normal buffered saline in a 13 mm by 

THE ROLE OF BIRDS IN ARBOVIRAL DISEASE SURVEILLANCE

IN HARRIS COUNTY AND THE CITY OF HOUSTON, TEXAS

Figure 1. Map of Harris County and the City of Houston, Texas, showing bird trap sites in 2015.
January – June 2017 3

THE ARMY MEDICAL DEPARTMENT JOURNAL

100 mm glass culture tube. Once blood collection was 

complete, the bird was banded and released.

The culture tubes were labeled with a unique identifying number and kept in a refrigerated cooler while in 

the field. Once the diluted blood samples arrived at the 

MVCD Avian Laboratory, they were placed in a refrigerated (4°C) centrifuge which ran for 15 minutes at 2,500 

rpm. The sera were extracted with disposable transfer 

pipettes into similarly labeled 2 ml cryovials, which were 

kept frozen until they were transferred to the University 

of Texas Medical Branch at Galveston (UTMB-G) or the 

MVCD Virology Laboratory for testing.

Most dead birds were reported to the Dead Bird Hotline, while a smaller number was reported to the HCPH 

website (https://secure.hcphes.org/MC/DeadBirdReport.html). Residents may also bring dead birds to the 

MVCD Avian Laboratory. Information collected on all 

birds included the contact information (name, address, 

and phone number) of the caller, location, condition of 

the bird, and time and date the bird was found. If the 

bird was found within 24 hours of the call and appeared 

reasonably fresh, the caller was instructed to double bag 

the bird and place it in the freezer or a bag of ice to preserve it until retrieved by a MVCD technician. In years 

with large numbers of dead bird calls, 18 additional drop 

off centers located throughout Harris County were activated. Once the bird was collected, it was identified and 

its condition evaluated. If the carcass was fresh and contained no maggots or ants, it was submitted for WNV/

SLE testing and assigned a unique identifying number.

Testing Procedures

Since 2002, the UTMB-G has been conducting the initial screening of live bird samples and confirmatory testing of dead birds. Sera samples were screened with the 

hemagluttination inhibition (HI) test, which detected 

the presence of viral antibodies, but could not differentiate between antibody subtypes. Since some subtypes, 

such as IgG, can persist for long periods of time, this test 

cannot be used to distinguish currently infected birds 

from those that were infected in previous years. West 

Nile virus and SLE are also structurally similar, causing 

HI antigens to cross-react. If the titer levels for one virus 

were 4 times or more titers higher than the other, the 

virus with the higher titer was considered the causative 

agent.12 If the causative virus was identified, the sample 

was considered positive solely for the causative virus. 

Samples in which the causative virus could not be determined were listed as WNV/SLE and analyzed separately.

To determine if the birds were recently infected, the 

MVCD Virology Laboratory preformed a second test, 

ie, the IgM antibody capture method, on all HI positive 

samples. This test does not cross-react and detects only 

IgM antibodies which are only produced during the initial stages of infection, and persist for about a month. A 

limitation of the IgM test is its potential for reacting to 

a naturally occurring antigen in the blood of some individual birds, resulting in a false positive rate of approximately 10%. Given these limitations, the most definitive 

results were from combining these tests. For example, a 

bird that tested both HI and IgM positive was definitely 

exposed and, most likely, that exposure was within the 

last month, indicating recent viral activity.

Dead birds were initially screened at the MVCD Avian 

Laboratory with vector and rapid analyte measurement 

platform (RAMP) test kits, both of which can also be 

used for mosquitoes. The vector test is a quick dipstick 

test that detects SLE and Eastern equine encephalitis 

(EEE) in addition to WNV. While the RAMP test takes 

longer, it is more accurate and provides quantitative results. For both tests, the bird’s throat was swabbed with 

a cotton-tipped applicator. For vector tests, the applicator was swirled in a 0.6 ml vial containing 600 μl of 

grinding solution for 10 seconds. A test strip was placed 

in the vial and left to mature for 15 minutes. Each virus 

had its own specific detection zone on the test strip. For 

RAMP testing, a separate applicator was swirled in a 

1.5 ml vial containing 1 ml of RAMP buffer solution for 

10 seconds. Then, 120 μl of the supernatant was transferred to a smaller 0.6 ml vial where it was mixed with 

a florescent dye. Finally, 70 μl was transferred to the 

sample well of the test cartridge and dried for 90 minutes. The dry cartridge was placed into a RAMP reader 

which provided a quantitative ratio in a range of less 

than 10 to more than 640. Results of 50 or greater were 

considered positive.

After testing, dead birds were labeled with weatherproof 

tags, double-bagged and placed in an ultra-low (-62°C) 

freezer until they were transferred to UTMB-G. Information on the tags included date, location, species, 

and MVCD Avian Laboratory number. If more than 25 

birds were collected during the week, only those that 

tested positive or species with historically high rates 

of WNV were transported to UTMB-G for tissue culture and WNV confirmation. During tissue culture, a 

small portion of brain was removed from each dead 

bird and was homogenized in approximately 2.0 ml of 

phosphate-buffered saline, pH 7.4, containing 10% fetal bovine serum. After centrifugation at 10,000 rpm for 

10 minutes, the supernatant was filtered (0.22 μm) and 

200 μL was inoculated into a flask culture of Vero cells. 

Vero cultures were maintained at 37°C for 14 days and 

observed daily for viral cytopathic effect (CPE). If CPE 
4 http://www.cs.amedd.army.mil/amedd_journal.aspx

was observed, the culture was confirmed as positive 

for WNV by complement-fixation test or by the vector 

test antigen assay done on the culture medium. Due to 

funding limitations, an alternative confirmation method 

consisting of a RAMP result of 100 or greater, corresponding to a UTMB-G confirmation rate of 90%, was 

implemented by MVCD Avian Laboratory from October 14, 2015 to December 31, 2015.

The capture and handling of wild birds was done in accordance with the US Federal Migratory Bird Treaty 

Act13 and required both federal and state permits. This 

work was conducted under Scientific Collecting Permit 

MB730179-0 from the US Fish and Wildlife Service, 

Federal Bird Banding Permit 09415 

from the US Geological Survey, and 

Scientific Research Permit SPR0816-179 from the Texas Parks and 

Wildlife Department.

Results

The MVCD Avian technicians collected a total of 1,345 sera samples 

and received 358 dead bird calls 

from 40 species of live bird species 

and 39 species of dead birds in 2015 

(Tables 1 and 2). Sera collections increased over 2014, improving upon 

the 50% increase over the number of 

samples collected in 2013 and surpassing all previous years since 2009. 

Of the dead birds called in, 

319 (89%) were collected and 

253 (79%) were fit for testing. 

Fifteen dead birds (5.9% of 

those tested) were confirmed 

positive for WNV, while 222 

(16.5%) sera samples were HI 

positive for WNV, 2 (0.15%) 

were HI positive for SLE 

and 10 (0.7%) were HI positive for either WNV or SLE. 

Additionally, 2 samples each 

were HI positive for EEE and 

Western equine encephalitis 

(WEE). Of the samples that 

were HI positive, 25 (10.6%) 

were IgM positive for WNV 

and 4 (1.6%) IgM positive for 

SLE. This represented about 

1.8% and 0.3% of all samples. 

The percentage of HI positive 

samples for WNV increased 

slightly over 2014, maintaining the generally elevated numbers recorded since 2012 

(Figure 2), while the percentage of IgM positives decreased by half or more (Figure 3). The percentage of 

positive dead birds also decreased from 2014 (Figure 4).

The percentage of WNV positives fluctuated throughout 

the year (Figure 5). The period of greatest activity was 

from July to September, 2015. There were no IgM positive samples in January. Between 5% and 9% of samples 

were IgM positive from February to June, 2015. This 

increased to 11% in July before peaking in August at 

28%. The first positive dead bird was found on July 2, 

2015. More followed, with all other positive dead birds, 

with one exception, occurring in July and August. The 

percentage of HI WNV positives 

peaked in September at 25%. The 

IgM WNV positives also remained 

high in September at above 20% before falling below 10% in October. 

There were no IgM positives in November and December. The last positive dead bird was found on November 6, 2015. The percentage of HI 

WNV positive samples was the only 

metric to remain elevated during the 

winter months, remaining at around 

15% for January, November, and December 2015. This is not surprising 

given the persistence of WNV antibodies and the high levels of WNV 

activity in 2012 and 2014.

THE ROLE OF BIRDS IN ARBOVIRAL DISEASE SURVEILLANCE

IN HARRIS COUNTY AND THE CITY OF HOUSTON, TEXAS

Year

Percentage Positive

20%

0%

5%

10%

15%

Figure 2. Percentage of live bird sera samples collected from Harris County and the City of 

Houston that tested HI positive for WNV, SLE, and WNV/SLE since 2002.

2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 2014 2015

WNV SLE WNV/SLE

Figure 3. Percentages of live bird seropositive samples collected from Harris County 

and the City of Houston, Texas that tested 

IgM positive for WNV and SLE in 2014 and 

2015. Percentage Positive

Year

20%

30%

0%

10%

25%

12.5%

2%

11%

SLE

WNV

2014 2015
January – June 2017 5

THE ARMY MEDICAL DEPARTMENT JOURNAL

Table 1. Number of live bird sera samples collected from Harris County and the City of Houston, Texas, that tested HI and IgM 

positive at the University of Texas Medical Branch, Galveston and the Mosquito and Vector Control Division in 2015.

Common Name Scientific Name Sera HI WNV HI SLE HI

WNV/SLE

HI EEE HI WEE IgM 

WNV

IgM

SLE

House Sparrow Passer domesticus 628 86 1 6 1 0 11 2

Blue Jay Cyanocitta cristata 158 46 0 1 0 0 10 2

Northern Cardinal Cardinalis cardinalis 117 54 0 1 0 0 2 0

Northern Mockingbird Mimus polyglottos 66 11 0 0 0 2 2 0

Mourning Dove Zenaida macroura 63 9 0 0 0 0 0 0

Myrtle Warbler Dendroica coronata 48 0 0 0 0 0 0 0

White-winged Dove Zenaida asiatica 37 3 0 1 0 0 0 0

Red-winged Blackbird Agelaius phoeniceus 34 0 1 0 0 0 0 0

European Starling Sturnus vulgaris 20 4 0 0 0 0 0 0

Common Grackle Quiscalus quiscula 18 0 0 0 0 0 0 0

Great-tailed Grackle Quiscalus mexicanus 18 2 0 0 0 0 0 0

Savannah Sparrow Passerculus sandwichensis 17 0 0 0 0 0 0 0

Brown-headed Cowbird Molothrus ater 15 1 0 0 0 0 0 0

Red-bellied Woodpecker Melanerpes carolinus 13 1 0 0 0 0 0 0

Chipping Sparrow Spizella passerina 12 0 0 0 0 0 0 0

American Robin Turdus migratorius 11 2 0 0 0 0 0 0

Nutmeg Manakin Lonchura punctulata 8 1 0 0 0 0 0 0

Carolina Wren Thryothorus ludovicianus 8 0 0 0 0 0 0 0

Tufted Titmouse Baeolophus bicolor 7 0 0 0 0 0 0 0

House Finch Carpodacus mexicanus 5 1 0 0 0 0 0 0

White-throated Sparrow Zonotrichia albicollis 5 0 0 0 0 0 0 0

Hermit Thrush Catharus guttatus 4 0 0 0 0 0 0 0

Inca Dove Scardafella inca 4 0 0 0 0 0 0 0

Yellow-bellied Sapsucker Sphyrapicus varius 4 0 0 0 0 0 0 0

Downy Woodpecker Picoides pubescens 3 0 0 0 0 0 0 0

Red-headed Woodpecker Melanerpes erythrocephalus 3 0 0 0 0 0 0 0

Bronzed Cowbird Molothrus aeneus 2 0 0 0 0 0 0 0

Brown Thrasher Toxostoma rufum 2 1 0 0 0 0 0 0

Eurasian Collared Dove Streptopelia decaocto 2 0 0 0 0 0 0 0

Loggerhead Shrike Lanius ludovicianus 2 0 0 0 0 0 0 0

Rock Dove Columba livia 2 0 0 0 0 0 0 0

Carolina Chickadee Parus carolinensis 1 0 0 0 0 0 0 0

Eastern Bluebird Sialia sialis 1 0 0 0 0 0 0 0

Eastern Phoebe Sayornis phoebe 1 0 0 0 0 0 0 0

Gray Catbird Dumetella carolinensis 1 0 0 1 0 0 0 0

Lincoln's Sparrow Melospiza lincolnii 1 0 0 0 0 0 0 0

Orange-crowned Warbler Vermivora celata 1 0 0 0 0 0 0 0

Purple Martin Progne subis 1 0 0 0 1 0 0 0

Rose-breasted Grosbeak Pheucticus ludovicianus 1 0 0 0 0 0 0 0

White-eyed Vireo Vireo griseus 1 0 0 0 0 0 0 0

(40 Bird Species) Totals 1,345 222 2 10 2 2 25 4

HI indicates hemagluttination inhibition. SLE indicates St Louis encephalitis. WEE indicates Western equine encephalitis.

WNV indicates West Nile virus. EEE indicates Eastern equine encephalitis. IgM indicates immunoglobulin M

Both HI and IgM results for WNV differed by age. After-hatch year birds had a HI seropositive rate more than 

double that of hatch year birds, while hatch year birds 

that were HI positive for WNV were twice as likely to be 

IgM positive (Table 3). This is consistent with after-hatch 

year birds maintaining protective immunity after being 

exposed to WNV within the previous 2 to 3 years. When 

disease prevalence in hatch year and after-hatch year 

birds was plotted by month, the differences were more 

pronounced (Figure 6). After-hatch year birds accounted 

for the majority of all positives until July, when hatch 

year IgM prevalence increased dramatically. After-hatch 
6 http://www.cs.amedd.army.mil/amedd_journal.aspx

THE ROLE OF BIRDS IN ARBOVIRAL DISEASE SURVEILLANCE

IN HARRIS COUNTY AND THE CITY OF HOUSTON, TEXAS

Table 2. Number of dead birds collected from Harris County and the City of Houston, Texas, that were tested 

and confirmed positive at the University of Texas Medical Branch, Galveston, and the Mosquito and Vector 

Control Division in 2015.

Common Name Scientific Name Tested Vector Test

Positive

RAMP

Positive

Confirmed

Positive

Percentage

Confirmed

White-winged Dove Zenaida asiatica 61 0 0 0 0.0%

Blue Jay Cyanocitta cristata 30 9 9 11 36.7%

European Starling Sturnus vulgaris 22 0 1 0 0.0%

Blue-winged Teal Anas discors 19 0 0 0 0.0%

House Sparrow Passer domesticus 18 0 0 0 0.0%

Great-tailed Grackle Quiscalus mexicanus 17 1 1 1* 5.9%

Mourning Dove Zenaida macroura 14 0 0 0 0.0%

Northern Mockingbird Mimus polyglottos 8 0 1 1 12.5%

American Robin Turdus migratorius 7 1 0 0 0.0%

Cedar Waxwing Bombycilla cedrorum 6 0 0 0 0.0%

Common Grackle Quiscalus quiscula 6 0 0 0 0.0%

Rock Dove Columba livia 4 0 0 0 0.0%

Yellow-crowned Night-heron Nycticorax violacea 4 0 1 0 0.0%

Cooper's Hawk Accipter cooperii 3 0 0 1 33.3%

Eastern Screech-owl Otus asio 3 0 0 0 0.0%

Northern Cardinal Cardinalis cardinalis 3 0 1 1 33.3%

Brown-headed Cowbird Molothrus ater 2 0 1 0 0.0%

Carolina Wren Thryothorus ludovicianus 2 0 0 0 0.0%

Eurasian Collared Dove Streptopelia decaocto 2 0 0 0 0.0%

Hermit Thrush Catharus guttatus 2 0 0 0 0.0%

Purple Martin Progne subis 2 0 0 0 0.0%

American Coot Fulica americana 1 0 0 0 0.0%

American Crow Corvus brachyrhynchos 1 0 0 0 0.0%

Baltimore Oriole Icterus galbula galbula 1 0 0 0 0.0%

Brown Pelican Pelecanus occidentalis 1 0 0 0 0.0%

Chipping Sparrow Spizella passerina 1 0 0 0 0.0%

Laughing Gull Larus atricilla 1 0 0 0 0.0%

Lesser Snow Goose Chen caerulescens 1 0 0 0 0.0%

Mallard Anus platyrynchos 1 0 0 0 0.0%

Myrtle Warbler Dendroica coronata 1 0 0 0 0.0%

Orange-crowned Warbler Vermivora celata 1 0 0 0 0.0%

Ovenbird Seiurus aurocapillus 1 0 0 0 0.0%

Red-eyed Vireo Vireo olivaceus 1 0 0 0 0.0%

Red-winged Blackbird Agelaius phoeniceus 1 0 0 0 0.0%

Sora Porzana carolina 1 0 0 0 0.0%

Tennessee Warbler Vermivora peregrina 1 0 0 0 0.0%

Veery Catharus fuscescens 1 0 0 0 0.0%

White-faced Ibis Plegadis chihi 1 0 0 0 0.0%

Yellow-breasted Chat Icteria virens 1 0 0 0 0.0%

(39 Bird Species) Totals 253 11 14 15 5.9%

*Includes bird confirmed in MVCD Avian Laboratory. RAMP indicates rapid analyte measurement platform.

year IgM rates did not undergo a similar increase until 

August. Hatch year HI rates were generally low, peaking 

at 21.5% in September, about 15% less than after-hatch 

year HI rates for the same month. There were no hatch 

year birds in January and February, and low sample sizes 

prevented the analysis of hatch year seropositive rates in 

March and April, which had one HI positive each.

Each species had a seropositive rate markedly different from the average. Northern Cardinals had the highest HI incidence rate of 46.1%, followed by Blue Jays 

at 29.1% (Figure 7). Of the HI positive samples, the 

majority were from House Sparrows (39%), Northern 

Cardinals (24%), and Blue Jays (21%) (Figure 8). The 

highest IgM incidence rates were in Blue Jays (21.3%), 

Mockingbirds (18.2%), and House Sparrows (11.8%), 

while Blue Jays and House Sparrows comprised the 

bulk of the IgM positive samples (84.0%). No HI or IgM 

positives were found in Myrtle Warblers, Red-Winged 

Blackbirds, Common Grackles, and Savanah Sparrows. 

Myrtle Warblers and Savanah Sparrows are migratory birds that only reside in the Houston area during 
January – June 2017 7

THE ARMY MEDICAL DEPARTMENT JOURNAL

the winter months. House Sparrows were by far the 

most frequently tested species, caught 4 times as much 

as Blue Jays, the next most commonly caught species, 

and comprised 46.7% of all samples tested. The next 4 

most commonly tested species were Blue Jays, Northern Cardinals, Northern Mockingbirds, and Mourning 

Doves, comprising 30.0% of all samples tested. 

Dead Blue Jays were also positive frequently, 

accounting for 73.3% of positive dead birds. 

All other dead positive bird species (Cooper’s 

Hawk, Northern Cardinal, Northern Mockingbird, and Great-tailed Grackle) had one positive sample each (Figure 9).

Positives were not uniformly distributed 

across Harris County. Parts of the Harris 

County that had high prevalence rates were 

the Spring-Cypress area in the northwest, 

Kingwood in the north, and western Houston. 

Figure 10 shows the distribution of HI positive 

results within Harris County for all viruses in Figure 5. Monthly WNV activity in live and dead birds collected from Harris 

County and the City of Houston, Texas, in 2015.

Month

Percentage Positive

0%

5%

30%

25%

20%

15%

10%

Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec

HI WNV IgM WNV Dead Birds

Table 3. Number and percentage of live birds collected in Harris County and the City of Houston, Texas, that tested HI 

and IgM positive in 2015.

2015 Total Sera

N

HI WNV

(%N)

HI SLE

(%N)

HI WNV/SLE

(%N)

HI EEE

(%N)

HI WEE

(%N)

IgM WNV+

(%N)

IgM SLE+

(%N)

All Birds 1,345 222 (16.5%) 2 (0.1%) 10 (0.9%) 2 (0.1%) 2 (0.1%) 25 (10.7%) 4 (1.7%)

Age

After Hatch Year 777 177 (22.8%) 1 (0.1%) 7 (0.9%) 1 (0.1%) 0 (0.0%) 16 (8.7%) 3 (1.6%)

Hatch Year 568 45 (7.9%) 1 (0.2%) 3 (0.5%) 1 (0.2%) 2 (0.3%) 9 (18.3%) 1 (2.0%)

Sex

Male 395 91 (23.0%) 1 (0.2%) 3 (0.7%) 0 (0.0%) 0 (0.0%) 7 (7.3%) 0 (0%)

Female 234 42 (17.8%) 1 (0.4%) 2 (0.8%) 1 (0.4%) 0 (0.0%) 0 (0.0%) 1 (2.2%)

Unknown 716 89 (12.4%) 0 (0.0%) 5 (0.7%) 1 (0.1%) 2 (0.2%) 18 (19.3%) 3 (3.2%)

HI indicates hemagluttination inhibition. SLE indicates St Louis encephalitis. WEE indicates Western equine encephalitis.

WNV indicates West Nile virus. EEE indicates Eastern equine encephalitis. IgM indicates immunoglobulin M.

Figure 4. Number and percentage of dead birds collected from Harris County and the City of Houston, Texas, that were tested and 

confirmed WNV positive since 2002.

Percentage WNV Positive

70%

60%

50%

40%

0%

10%

20%

30%

Number of Dead Birds

2500

2000

1500

1000

500

0

2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 2014 2015

57.2%

18.6%

24.5%

12.7%

14.1%

9.2%

0.3% 3.0% 1.9%

5.3%

32.5%

1.6%

23.5%

5.9%

Year

WNV Lab Tested % WNV Positive
8 http://www.cs.amedd.army.mil/amedd_journal.aspx

2015. The prevalence rate for WNV ranged 

from 0.0% to 28.6%, and areas with the highest percentages were in the center and southern part of Harris County. Only 6 samples 

tested positive for SLE, EEE, and WEE. Areas 411 and 425 tested positive for SLE, while 

EEE was detected in areas 820 and 904, and 

WEE was detected in 710 and 923, which was 

the first time it was detected since 2011.

Figure 11 shows the distribution of IgM results 

for WNV in Harris County. The percentage 

of IgM positive samples is the percentage of 

HI samples testing IgM positive and indicates 

the proportion of HI positives in each area that 

is due to a recent infection. The 

percentage of IgM positives in 

each area ranged from 0.0% to 

36.4%. The areas with the highest percentage of IgM positives 

were 820, 225, and 606. Area 606 

was the only one that had a high 

percentage of both IgM and HI 

positives.

Most dead birds were reported 

from the western loop and northwestern Harris County, near the 

towns of Spring and Cypress 

(Figure 12). Positive birds were 

clustered in Baytown and near 

the intersection of highways 610 

and 290. Five more birds were 

confirmed from outlying areas 

northwest of the Sam Houston 

Beltway and one from the town 

of Humble.

Comparisons to Previous Years

In 2015, the percentage of positive confirmed dead birds 

was low, while the percentage of HI WNV positive live 

birds was high, relative to their historical occurrences 

(Figure 13). This could be explained by the high prevalence of antibodies in the avian population providing 

protective immunity against WNV resulting in the avian low death rate, which is reasonably well supported by 

the historical data. However, high antibody levels did 

not prevent a high death rate in 2014.

In 2002 when WNV was first detected in Harris County, 

birds were particularly susceptible as none had been 

previously exposed. Although SLE antibodies provide 

some protection against WNV,6

 they were at a historically low rate of 1.9%. Mortality was high, particularly 

for American Crows (68.4%) and Blue Jays (66.9%). It 

took 4 years for antibody levels to peak in 2006 at 17.7%, 

double the highest annual SLE rate from 1975 to 2001 of 

8.4%. From 2006 to 2011, the mortality and seroprevalence rates decreased, until antibody levels reached 6% 

to 5% (Figure 13). In 2012, there was a sudden resurgence in WNV activity, with mortality rates reaching 

2003-2004 levels. Antibody levels jumped to 14.7% and 

remained high throughout 2015. Recent mortality rates 

were inconsistent; 1.6% in 2013 and 23.5% in 2014.

Program Expansion

The objectives were to expand and improve the MVCD 

Avian Surveillance Program in 2016. Trapping sites were 

continually evaluated and subjected to change if catch 

rates decreased. Scouting for new trapping locations, 

THE ROLE OF BIRDS IN ARBOVIRAL DISEASE SURVEILLANCE

IN HARRIS COUNTY AND THE CITY OF HOUSTON, TEXAS

Northern Cardinal

American Robin

Northern Mockingbird

Mourning Dove

House Sparrow

Great-tailed Grackle

White-winged Dove

Red-bellied Woodpecker

Brown-headed Cowbird

Other

Myrtle Warbler

Red-winged Blackbird

Common Grackle

Savannah Sparrow

Chipping Sparrow

Blue Jay

European Starling

0% 5% 10% 15% 20% 25% 30% 35% 40% 45% 50%

Figure 7. Percentages of live bird sera samples collected from Harris County and the City 

of Houston that tested HI positive for WNV in 2015.

Figure 6. Monthly WNV activity in live hatch year and after-hatch year birds 

collected from Harris County and the City of Houston, Texas, in 2015.

Month

Percentage Positive 10%

5%

0%

-5%

20%

40%

35%

30%

25%

15%

Hatch Year HI

Hatch Year IgM

After-hatch Year HI

After-hatch Year IgM

Jan Feb Mar Apr May Jun Jul Aug Sep Oct Nov Dec
January – June 2017 9

THE ARMY MEDICAL DEPARTMENT JOURNAL

Figure 8. Species distribution of HI positive live bird samples 

collected from Harris County and the City of Houston, Texas, in 

2015.

Northern

Cardinal

24.3%

House

Sparrow

38.7%

Other

7.2%

Blue Jay

20.7%

Northern

Mockingbird

5%

Mourning Dove

4.1%

Northern

Cardinal

6.6%

Great-tailed

Grackle 39%

Cooper’s Hawk 6.6%

Blue Jay

73.3%

Northern Mockingbird

6.6%

Figure 9. Species distribution of WNV dead birds collected from 

Harris County and the City of Houston, Texas, in 2015.

Figure 10. Map of Harris County and the City of Houston, Texas, showing the geographical

distribution of HI test results for WNV, SLE, EEE, and WEE in 2015.
10 http://www.cs.amedd.army.mil/amedd_journal.aspx

particularly in areas of high virus activity or low current coverage, were conducted as time permitted. Testing was expanded to include Highlands J virus (HJV), 

an alphavirus in the WEE complex. Although HJV is 

not thought to cause illness in humans, it can react with 

antigens for WEE similarly to WNV and SLE. Testing 

for both will allow for definitive identifications of WEE. 

Five sera samples have already tested HI positive for 

HJV in 2016.

Testing for non-arboviral viruses, such as Newcastle 

disease virus (NDV) and highly pathogenic avian influenza (HPAI) can also be considered. The UTMB-G has 

frequently recovered NDV, which is classified as a select 

agent by the US Department of Agriculture in pigeons 

and doves collected in Harris County and the City of 

Houston.14 Although HPAI is not a vector-borne disease, 

it is a high-profile, economically devastating, and potentially life-threatening disease that uses avian species as 

the main host. In 2015, an HPAI outbreak occurred in at 

least 15 states (3 declaring states of emergency) resulting in the destruction of 49.5 million birds and at least 

$500 million spent in emergency funds.15-17 The HPAI 

was also found in wild birds in Kansas and Missouri,18

states that are part of the same migratory flyway that 

includes important wintering grounds in the Houston 

area. The HPAI may persist in wild birds and resurface 

from time to time in the United States as it has in Asia. 

Indeed, the 2015 outbreak was linked to incidences in 

British Columbia and the Pacific Northwest in 2014.19

Another incidence was confirmed on January 15, 2016, 

in Indiana where 43,000 birds were destroyed as part of 

the containment effort.20

We plan to test for Avian Flu in 2017. Test kits are available from several commercial manufactures for use by 

veterinarians. Further testing by UTMB-G will be needed to confirm any positive samples as highly pathogenic.

THE ROLE OF BIRDS IN ARBOVIRAL DISEASE SURVEILLANCE

IN HARRIS COUNTY AND THE CITY OF HOUSTON, TEXAS

Figure 11. Map of Harris County and the City of Houston, Texas, showing the geographical distribution of IgM test results for WNV in 2015.
January – June 2017 11

THE ARMY MEDICAL DEPARTMENT JOURNAL

Conclusion

Avian surveillance is an integral part of disease-based 

integrated mosquito and vector management programs. 

Overall, an average level of Avian WNV activity was indicated in 2015, and low levels of EEE and SLE in Harris 

County and the City of Houston. However, WEE was 

last detected in 2011. Most infections were in residential 

birds. If viral activity conforms to SIR models, Harris 

County should be entering a low point in the WNV cycle. Initial results showed WNV activity was low in 2016, 

with only one confirmed dead bird. Low levels of viral 

activity will eventually diminish avian arboviral protection, inviting another outbreak. Continued surveillance 

is needed to determine when this will likely occur.

References

1. Rappole JH, Hubalek Z. Migratory birds and West 

Nile virus. J Appl Microbiol. 2003;94(suppl):47s-58s.

2. McKee EM. West Nile Seroreversion and the Influence of Herd Immunity on Disease Risk in a LongTerm Study of Free-Ranging Birds [master’s thesis]. 

University Park, IL; Governors State University; 

2012. Available at: http://opus.govst.edu/theses/9/. 

Accessed December 14, 2016.

3. Nemeth NM, Oesterlem PT, Bowen RA. Humoral 

immunity to West Nile virus is long-lasting and 

protective in the House Sparrow (Passer domesticus). Am J Trop Med Hyg. 2009;80(5):864-869.

4. Nash D, Mostashari F, Fine A, et al. The outbreak 

of West Nile virus infection in the New York City 

area in 1999. N Engl J Med. 2001;344:1807-1814.

5. Koenig WD, Hochachka WM, Zuckerberg 

B, Dickinson JL. Ecological determinants of 

American crow mortality due to West Nile virus 

during its North American sweep. Oecologia. 

2010;163:903-909.

Figure 12. Map of Harris County and the City of Houston, Texas, showing the geographical distribution of dead birds reported, tested, and 

confirmed WNV positive in 2015.
12 http://www.cs.amedd.army.mil/amedd_journal.aspx

6. Fang Y, Reisen WK. Previous infection with West 

Nile or St. Louis encephalitis viruses provides 

cross protection during reinfection in house finches. Am J Trop Med Hyg. 2006;75(3):480-485.

7. Anderson RM, May RM. Infectious diseases of 

Humans: Dynamics and Control. New York, NY: 

Oxford University Press; 1991.

8. Kwan JL, Kluh S, Reisen WK. Antecedent avian 

immunity lifts tangential transmission of West 

Nile virus to humans. PLoS One. 2012;7:e34127.

9. Dunn JL. National Geographic Field Guide to the 

Birds of North America, 4th ed. Washington, DC: 

National Geographic Society. 2002.

10. Sibley DA. The Sibley Field Guide to Birds of 

Eastern North America. New York, NY: Alfred A. 

Knopf, Inc; 2003.

11. Pyle P. Identification Guide to North American 

Birds, Part 1. Bolinas, CA: Slate Creek Press; 1997.

12. Casals J. The arthropod-borne group of animal viruses. Trans N Y Acad Sci. 1957;19(3):219-235.

13. Migratory Bird Treaty Act of 1918, 16 USC 

§703-712; Ch 128; 40 Stat. 755 (July 13, 1918). 

Available at: http://law2.house.gov/view.xhtml?

req=granuleid%3AUSC-prelim-title16-chapter7-su

bchapter2&saved=%7CMTYgdXNj%7CdHJlZXN

vcnQ=%7CdHJ1ZQ==%7C5302%7Ctrue%7Cprel

im&edition=prelim. Accessed December 14, 2016.

14. Kim LM, Guzman H, Tesh RB, Bueno R, Dennett 

JA, Alonso CL. Biological and phylogenetic characterization of pigeon paramyxovirus serotype-1 

circulating in North American pigeons and doves. 

J Clin Microbiol. 2008;46:3303-3310.

15. McKenna M. The avian flu epidemic: massive impact, uncertain future [The National Geographic 

Society Web site]. May 7, 2015. Available at: http://

phenomena.nationalgeographic.com/2015/05/07/

bird-flu-1/. Accessed December 14, 2016.

16. McKenna M. Bird flu cost the US $3.3 billion and 

worse could be coming [The National Geographic 

Society Web site]. June 15, 2015. Available at: http://

phenomena.nationalgeographic.com/2015/07/15/

bird-flu-2/. Accessed December 14, 2016.

17. Lowe P, Boden S. Avian flu outbreak takes poultry 

producers into uncharted territory [NPR Web site]. 

May 21, 2015. Available at: http://www.npr.org/

sections/thesalt/2015/05/21/408306843/avian-fluoutbreak-takes-poultry-producers-into-unchartedterritory. Accessed September 19, 2016.

18. US Department of Agriculture. USDA list of confirmed US highly pathogenic avian influenza wild 

bird cases in the United States. Updated September 

4, 2015. Available at: www.CDC.gov/flu/avianflu/

h5/.

19. US Geological Survey Instructional Memorandum 

No: IM OA 2014-05. Detection of Highly Pathogenic Avian Influenza Viruses H5N2 and H5N8 in 

wild birds of the United States. July 7, 2014.

20. US Department of Agriculture. Confirmed Avian 

Influenza Detections – 2016. Available at: https://

www.aphis.usda.gov/aphis/ourfocus/animalhealth/

animal-disease-information/avian-influenza-disease/sa_detections_by_states/ai-2016-map. Accessed September 19, 2016.

Authors

Ms Wilkerson is the Avian Biologist at the Mosquito and 

Vector Control Division, Harris County Public Health, 

Houston, Texas.

Mr Reyna Nava is a Medical Entomologist and the Technical Operations Manager of the Mosquito and Vector 

Control Division, Harris County Public Health, Houston, 

Texas.

Ms Freeman is the Virologist at the Mosquito and Vector 

Control Division, Harris County Public Health, Houston, 

Texas.

Ms Travassos da Rosa is a Virologist at the Department of Pathology, University of Texas Medical Branch, 

Galveston, Texas.

Ms Guzman is a Virologist at the Department of Pathology, University of Texas Medical Branch, Galveston, 

Texas.

Dr Tesh is a Professor at the Department of Pathology, 

University of Texas Medical Branch, Galveston, Texas.

Dr Debboun is the Director of the Mosquito and Vector 

Control Division, Harris County Public Health, Houston, 

Texas.

THE ROLE OF BIRDS IN ARBOVIRAL DISEASE SURVEILLANCE

IN HARRIS COUNTY AND THE CITY OF HOUSTON, TEXAS

Figure 13. Percentage of 

dead birds confirmed positive and live birds that tested HI positive for WNV from 

Harris County and the City of 

Houston, Texas, since 2002.

Percentage Positive

0%

20%

60%

50%

40%

30%

10%

2002 2003 2004 2005 2006 2007 2008 2009 2010 2011 2012 2013 2014 2015

Dead Birds Live Birds
January – June 2017 13

Aedes aegypti (L.), also known as the “yellow fever mosquito,” is an important vector of viral diseases such as 

yellow fever, dengue, Chikungunya, and, most recently, 

Zika virus.1,2 Despite current vector control efforts, the 

incidence and geographical expansion of these pathogens continue to increase at an alarming rate. For example, the incidence of dengue has increased 30 fold 

over the last 50 years.3

 The World Health Organization 

(WHO) estimates 50-100 million cases occur annually 

in over 100 endemic countries, putting almost half of 

the world’s population at risk. More than 70% of the 

population at risk for dengue worldwide live in member 

states of the WHO southeast Asia and western Pacific 

region.3

 As a result, the Department of Defense (DoD) 

personnel and family members stationed in these regions are at risk.

Vector control remains the primary defense against 

vector-borne diseases. Standard preventive deterrence 

against vector-borne diseases incorporates integrated 

pest management (IPM) with personal protective measures (PPMs). The IPM approach to pest control integrates cultural, biological, physical, and chemical methods which minimize health and environmental risk. Often in situations when the threat of vector-borne disease 

transmission is high, direct suppression of vectors using 

pesticides is favored over nonchemical control methods. 

However, the repeated failure of conventional insecticides to control vector species in some areas continues 

to put Soldiers at risk. Furthermore, the failure to control vector populations may result in additional chemical 

applications that potentially contribute to other vector 

control problems such as pesticide resistance, unintended exposure to nontarget organisms, and environmental 

pollution.

The PPM standards mandated by the military include 

use of a topical repellent, N, N-diethyl-3-methylbenzamide (DEET) or 1-piperidinecarboxylic acid 2-

(2-hydroxyethyl)-1-methylpro-pylester) (Picaridin), prop

er wear of a permethrin-treated uniform, and the use 

of a bed net.4

 Additionally, malaria prophylaxis is prescribed when service members are deployed in endemic 

areas. However, compliance with PPM standards remains a problem among service members.5-7 Sanders et 

al8

 reported that responders in a 2004 study of personnel returning from Iraq and Afghanistan indicated that 

a majority (51%) did not use DEET, even though most 

knew that it was readily available. A report on the use of 

PPM among confirmed malaria cases in 2012 revealed 

that compliance with 4 preventive measures (use of 

DEET, proper uniform wear, adherence to prophylaxis, 

Laboratory and Semi-field Evaluations

 of Two (Transfluthrin) Spatial Repellent

 Devices Against Aedes aegypti (L.)

 (Diptera: Culicidae)

MAJ Lee P. McPhatter, MS, USA CPT Erica J. Lindroth, MS, USA

Paula D. Mischler, PhD Alec G. Richardson, PhD

Meiling Z. Webb, PhD Mustapha Debboun, PhD, BCE

Kamal Chauhan, PhD

Abstract

Two transfluthrin-based spatial repellent products (Raid Dual Action Insect Repellent and Home Freshener and 

Raid Shield (currently not commercially available), SC Johnson, Racine WI) were evaluated for spatial repellent 

effects against female Aedes aegypti (L.) mosquitoes under laboratory (wind tunnel) and semi-field (outdoor 

enclosure) conditions. The placement of either product in the wind tunnel significantly reduced host-seeking 

behaviors. The mean baseline (control) landing counts for the Raid Dual Action and Raid Shield were reduced 

by 95% and 74% respectively. Mean probing counts for the Raid Dual Action were reduced by 95%, while the 

probing counts for the Raid Shield were decreased by 69%. Baseline blood-feeding success was significantly 

reduced for both treatments: Raid Dual Action (100%) and Raid Shield (96%). Semi-field evaluations were conducted in outdoor enclosures at the Navy Entomology Center of Excellence, Jacksonville, Florida. A moderate 

reduction in mosquito entry into military style tents resulted when either product was placed near the tent opening. The Raid Shield reduced mosquito entry into tents by 88%, while the Dual Action decreased entry by 66%.
14 http://www.cs.amedd.army.mil/amedd_journal.aspx

and postdeployment antirelapse therapy) was 0%.9 Even 

when compliance with PPM is high, service members 

are still exposed to vectors during physical training or 

while off duty. Additional control methods are needed to 

augment the DoD’s current vector control strategy.

Spatial Repellents

Recently, area (spatial) repellents have received interest as a novel system for vector control. Spatial repellents work by releasing chemicals into the air to reduce 

mosquito entry into treated spaces and to inhibit bloodfeeding behavior. An additional benefit of this type of 

behavioral modification is a delayed or diminished development in the emergence of insecticide resistance.10

Several types of commercial spatial repellent products 

have been reported in the literature: impregnated plastic 

or paper strips,11 coils,12-14 candles,15-17 fan emanators,18-21

heat-generating devices,20-22 and microdispensers.23

Impregnated Plastic

Plastic or other substrates impregnated with fluorinated 

pyrethroids (metofluthrin and transfluthrin) can passively deliver spatial repellents without external energy 

input. Yayo et al11 investigated the effect of Rambo Insecticide Paper (0.45% transfluthrin) on indoor resting 

densities and biting rates of 2 important malaria vectors (Anopheles funestus Giles and An. gambiae s.l.) at a 

peri-urban field site in Nigeria. A significant reduction in 

both indoor resting density (91%) and biting rate (86%) 

was reported. Passive emanators may be preferred by 

the DoD, as these devices would be most likely to meet 

logistical and safety requirements.

Mosquito Coils

Mosquito coils are the most commonly used household 

insecticidal product in the world.12 Avicor et al13 conducted a laboratory evaluation of 3 commercial coil 

products for protection efficacy against An. gambiae 

Giles in southern Ghana. All 3 coils (FISH M (0.005% 

metofluthrin)) 86%, Shieldtox (0.15% d-trans allethrin) 

74%, and Tesco Value (0.3% allethrin) 72% showed high 

knockdown efficacy. Hill et al12 demonstrated the effectiveness of using the Transfluthrin mosquito coil (Raid, 

SC Johnson, Racine, WI) in households to reduce malaria transmission. The Transfluthrin mosquito coil provided 77% protection against Plasmodium falciparum 

Welch. Msangi et al14 assessed feeding inhibition and 

repellency of 3 pyrethroid brands (Kiboko, Total, and 

Risasi) of mosquito coils in experimental huts in northern Tanzania. Induced exophily was high for Culex 

quinquefasciatus Say (95%), but was low in An. gambiae (61%) for all products tested. Feeding inhibition was 

significantly reduced for both species: Cx. quinquefasciatus (97%) and An. gambiae (88%).

Fan Emanators

Fan emanators that vaporize metofluthrin are marketed 

as promising tools with long-lasting spatial protection 

against mosquitoes. Various studies have evaluated the 

efficacy of the Off! Clip-On device (SC Johnson). Xue 

et al18 evaluated this device in the field in northeastern 

Florida, which proved to be 70% and 79% effective at 

repelling Aedes albopictus Skuse and Ae. taeniorhynchus (Wiedemann) from human test subjects. In a semifield study in Israel, the device reduced biting on the 

arms of volunteers against Ae. albopictus by 96% and 

Cx. pipiens L. by 95%.19 Lloyd et al20 reported a 64% 

reduction in Ae. albopictus collected from Biogents 

Sentinel (BGS) traps located near the repellent device. 

The Off! Clip-On device has also been reported to have 

limited to no effect on certain species. In a semi-field 

study conducted by Dame et al,21 the device failed to 

reduce the populations of An. quadrimaculatus Say and 

Psorophora columbiae (Dyar and Knab) collected in 

surrogate traps. Reduction rates for this study ranged 

from 8% to 16%.

Heat Generating Devices

The ThermaCell (Thermacell Repellents, Bedford, MA) 

spatial repellent device (active ingredient allethrin), is 

one of the most popular heat generating devices on 

the market. Collier et al22 conducted a study in Louisiana using several spatial repellent devices (Dragon 

fly/Mosquito Cognito, ThermaCell, SC Johnson OFF! 

Mosquito Lantern) and systems to determine their effect on backyard mosquito population levels, in which 

the ThermaCell device was the most effective device in 

reducing mosquito populations (Cx. salinarius Coquillett, Cx. quinquefasciatus, and Ae. vexans (Meigen)). In 

a field study conducted in Jacksonville Florida,20 the 

number of Ae. albopictus repelled by the ThermaCell 

was significantly higher than the other spatial repellents 

evaluated (Lentek Bite Shield and Bug Button Mosquito 

Eliminator).

The objectives for this study were (1) to evaluate the efficacy of two commercially available transfluthrin spatial 

repellents for inhibiting Ae. aegypti (L.) host-seeking 

behaviors (anemotaxis, landing, and blood feeding) 

under laboratory conditions, and (2) determine if these 

products were effective in reducing the number of hostseeking Ae. aegypti from entering a baited militarystyle tent in a controlled semi-field environment.

Materials and Methods

Mosquitoes

Aedes aegypti eggs (Liverpool and Orlando strain) 

were obtained from the US Department of Agriculture 

(USDA) Agricultural Research Service (ARS) Center 

LABORATORY AND SEMI-FIELD EVALUATIONS OF TWO (TRANSFLUTHRIN) SPATIAL REPELLENT DEVICES

AGAINST AEDES AEGYPTI (L.) (DIPTERA: CULICIDAE)
January – June 2017 15

THE ARMY MEDICAL DEPARTMENT JOURNAL

for Medical, Agricultural, and Veterinary Entomology, 

Gainesville, Florida. They were reared in an insectary 

using a 12:12 hour (light/dark) photoperiod at 26oC and 

35% to 50% relative humidity (RH). Larvae were fed 

ground tropical fish flakes (Tetra Werke, Melle, Germany). Adults were held in screened, 3.79 liter plastic buckets, and fed with a cotton pad moistened with 10% aqueous sucrose solution. Adult female mosquitoes were 5 to 

10 days old and incubated in plastic buckets along with a 

water-moistened pad without sugar for 24 hours before 

they were used in bioassays.

Spatial Repellent Devices Evaluated

Two transfluthrin-based spatial repellent (SC Johnson 

Company) products: Raid Dual Action Insect Repellent 

and Home Freshener, and Raid Shield were evaluated in 

this study (Figure 1).

In 2012, SC Johnson, Cornell University’s Center for 

Sustainable Global Enterprise, and the Bill and Melinda 

Gates Foundation created the WOW, which is a business concept that creates access to pest control products 

that can help prevent malaria in at-risk populations at 

the “base of the pyramid,” as well as home-cleaning and 

personal care products valued by rural consumers. The 

Raid Dual Action Insect Repellent and Home Freshener 

is one of 6 products developed from the 

WOW program in Ghana. The product consists of a spray bottle (0.40% transfluthrin) 

and a colorful plastic poster decorated with 

a well-known Ghanaian adinkra symbol.

Raid Shield is a product developed for research purposes (not commercially available) by the SC Johnson Company. It is a 

clear plastic strip that is impregnated with 

transfluthrin (1%). Once the plastic strip is 

exposed to air, transfluthrin is slowly released into the immediate area.

Laboratory Evaluations (Wind Tunnel)

Laboratory evaluations were conducted in a 3.0 m x 61 

cm x 60 cm wind tunnel (Figure 2A) at the USDA-ARS 

Invasive Insect Biocontrol and Behavior Laboratory 

in Beltsville, Maryland. With a few minor deviations, 

the wind tunnel was set up in accordance with Klun et 

al.24 Air temperature in the chamber was controlled by 

a window air conditioner and baseboard electric heaters and maintained at 25oC±4oC in the tunnel. The RH 

inside the chamber was maintained at 25% to 35% using 

a misting humidifier. The upwind end of the tunnel was 

connected to the air conditioning chamber by a cowling 

that housed a 61 cm diameter fan driven by a rheostatcontrolled Dayton Electric Model 2Z846A motor that 

pushed conditioned air through an air filter into the tunnel. Air flow within the tunnel was monitored using an 

air flow gauge, while temperature and relative humidity 

was recorded using a HOBO data logger (Onset Computer Corp, Bourne, MA).

The wind tunnel was fitted with a 6-well mosquito feeding reservoir (Figure 2B) that was positioned at the upwind end of the tunnel parallel to tunnel airflow. The 

reservoir was placed on a stand made of 0.5 cm thick 

Plexiglas with a 33.8 cm x 15.4 cm base and a 29.8 cm x 

7.0 cm (length x width) upper shelf with 1.2 cm high edges at each end of the shelf to secure the reservoir on the 

A B

Figure 1. The SC Johnson spatial repellents tested in this study: (A) Raid Dual 

Action Insect Repellent and Home Freshener; (B) Raid Shield.

Figure 2. (A) Wind tunnel at the Invasive Insect Biocontrol & Behavior Laboratory, USDA-ARS, Beltsville, MD. (B) Mosquito feeding 

reservoir.

A B
16 http://www.cs.amedd.army.mil/amedd_journal.aspx

shelf. Positioned on the shelf, the top surface of the reservoir was 18 cm above the tunnel floor, at the center of the 

tunnel, and 47 cm away from the upwind tunnel screen. 

In order to maintain a constant temperature (38oC) for 

the blood contained in the reservoir, it was modified and 

connected to a water bath circulator (Lauda E100, Wobser GMBH and Co, Konigshofell, Germany). A breathdelivery tube (2.5 m x 22 mm inner diameter; Tri-Anim, 

Sylmar, CA) ran from outside of the upwind end of the 

tunnel to breath exhaust position inside the tunnel 20 cm 

above the tunnel floor, at the tunnel center, and 20 cm 

from the upwind end of the tunnel.

Prior to each test, the upper surface of the feeding reservoir was coated with a thin layer of high-vacuum silicone grease (Dow Corning Corp, Midland, MI), and the 

wells were filled (approximately 6 ml capacity) with human blood from the Walter Reed Army Institute of Research (WRAIR), Silver Spring, MD, to which Adenosine triphosphate (ATP) was added on the day of testing 

to obtain a concentration of 10–3 mmol/L ATP. The filled 

cells were covered with an Edicol collagen membrane 

(G Street Fabrics, Rockville, MD). During treatment 

evaluations, the treatment device was suspended from 

the wind tunnel ceiling facing downwind and positioned 

38 cm in front of the feeding reservoir.

Trials for each spatial repellent device were replicated 5 

times. Each trial consisted of one control test followed 

by 5 consecutive treatment tests. The purpose of the 

control evaluation was to establish a baseline for mosquito activity under normal conditions (untreated) and 

to confirm the absence of pesticide contamination in the 

wind tunnel. A test began by placing a mosquito release 

canister containing 20 female Ae. aegypti (Liverpoolstrain) inside the wind tunnel. Subsequently, human 

breath was expired from the breath-delivery tube for 5 

seconds and the mosquitoes were released and allowed 

to freely fly inside the wind tunnel for 10 minutes. During this time period an observer recorded the frequency 

of probing and landing occurring on the mosquito feeding reservoir. Upon completion of the 10-minute period, 

the mosquitos were visually examined for evidence of 

knockdown from chemical exposure and feeding status 

(blood-fed or unfed). Mosquitoes were removed from 

the chamber using a vacuum aspirator.

Semi-field Evaluations (Enclosures)

Semi-field evaluations were conducted using 2 outdoor 

enclosures (Figure 3A), located at the Navy Entomology 

Center of Excellence in Jacksonville, Florida. Each enclosure consisted of a metal frame that measured 6.1 m 

wide x 10.7 m long x 3.4 m high. The enclosure was completely covered with mosquito screen and had a concrete 

base. Several potted plants were placed in the interior of 

each enclosure to mimic the natural outdoor habitat. A 

single military style tent (3-person) was placed at the far 

end of the enclosure, opposite the entrance. Each tent 

housed a BGS 2.0 trap baited with dry ice and BGS proprietary lure. Temperature, relative humidity, and wind 

speed were measured throughout the evaluation using a 

HOBO data logger.

Enclosure experiments were conducted simultaneously 

in the 2 enclosures. Prior to each trial, one of the enclosures was randomly selected as the treatment and 

the other as the control. Inside the treatment enclosure, 

the spatial repellent device was placed on the outside of 

the tent near the upper door opening (Figure 3B). Trials 

were conducted in the morning (8 am to 11 am) which 

corresponded with the peak host-seeking period of Ae. 

aegypti. Two hundred female Ae. aegypti (Orlandostrain) were simultaneously released into each enclosure 

and allowed to move freely for one hour. At the end of 

the trial, the mosquitoes inside the tents were collected 

from the BGS traps and vacuumed out using an aspirator.

LABORATORY AND SEMI-FIELD EVALUATIONS OF TWO (TRANSFLUTHRIN) SPATIAL REPELLENT DEVICES

AGAINST AEDES AEGYPTI (L.) (DIPTERA: CULICIDAE)

A

B

Figure 3. (A) A semi-field enclosure used in this study. (B) A commercial tent used in this study.
January – June 2017 17

THE ARMY MEDICAL DEPARTMENT JOURNAL

Statistical Analysis

Laboratory (Wind Tunnel) Study

Statistical 2-way hypothesis tests were conducted at the 

95% confidence level (α=0.05) to assess differences 

in landing, probe, blood-fed, and knock down counts 

among the 2 products (Raid Dual Action and Raid 

Shield) and treatments (Control and treatment).

Semi-field Study

Statistical 3-way and 2-way hypothesis tests were conducted at the 95% confidence level (α=0.05) to assess 

differences in mosquito counts (inside tent and BGS 

traps) between the 2 products (Raid Dual Action and 

Raid Shield), between the treatment and control, and 

between the 2 enclosures. Preliminary goodness-of-fit 

testing using the Kolmogorov-Smirnov test for normality25 and the Bartlett test for homoscedacity (homogeneity of variances)26,27 indicated that the data were nonnormal and non-homoscedastic. Thus, the nonparametric Kruskal-Wallis test28 was used instead of the parametric ANOVA test for the statistical analysis. Post hoc 

Tukey multiple-comparison,29 Newman-Keuls, Duncan 

multiple-range,30,31 and Scheffe multiple-contrast32 tests 

were evaluated based on an optimization analysis, and 

the optimal model was used to identify the specific pairwise combinations that are significantly different from 

each other and contribute to the overall variance source. 

Interaction effects of air temperature and humidity on 

mosquito counts were evaluated by regression analysis. 

All statistical analysis was performed using Intel Visual 

Fortran Compiler XE 2013 (Intel Corporation, Santa 

Clara, CA).

Results

Laboratory (Wind Tunnel)

Both products were effective in reducing host-seeking 

activity of Ae. aegypti in the wind tunnel. Figure 4A 

shows the treatment effect of both products on the mean 

landing counts of the mosquitoes. The mean baseline 

(control) landing counts for the Raid Dual Action and 

Raid Shield were reduced by 95% (P<.006) and 74% 

(P<.006) respectively. Mean probing counts for the Raid 

Dual Action were reduced by 95% (P<.02), while the 

probing counts for the Raid Shield were decreased by 

69% (P<.02) (Figure 4B). Baseline blood-feeding success was significantly reduced (P<.002) for both treatments: Raid Dual Action (100%) and Raid Shield (96%) 

(Figure 4C). Figures 5 and 6 show the time analysis (50 

minutes) for the mean counts of landing, probing, and 

blood feeding for each treatment. Maximum reduction 

rates occurred during the first 30 minutes. After 30 minutes, reduction rates dropped below 50%. Blood-feeding 

reduction rates remained above 85% for the duration of 

a

a

b

b

0

3

5

7

1

2

4

6

Raid Dual Action Raid Shield

Control Treatment

Mean Number of Blood-Fed Mosquitoes

C

10

0

20

30

40

50

60

70

a

a

b

b

Raid Dual Action Raid Shield

Control Treatment

Mean Number of Mosquito Probe Counts

B

20

0

40

60

80

100

120

140

160

Mean Number of Mosquito Landings

Raid Dual Action Raid Shield

a

a

b

b

A

Control Treatment

Figure 4. Treatment effects of both products on host-seeking activity: (A) Mean (±SE) mosquito landing counts; (B) Mean (±SE) 

number of mosquito probes; (C) Mean (±SE) number of bloodfed mosquitoes. Note: Bars tagged with the same letter are not 

significantly different.
18 http://www.cs.amedd.army.mil/amedd_journal.aspx

LABORATORY AND SEMI-FIELD EVALUATIONS OF TWO (TRANSFLUTHRIN) SPATIAL REPELLENT DEVICES

AGAINST AEDES AEGYPTI (L.) (DIPTERA: CULICIDAE)

Time (minutes)

Control Treatment

Mean Number of Blood-Fed Mosquitoes

Raid Dual Action on Blood Feeding

3

5

1

0

2

4

6

10 20 30 40 50

C

Raid Dual Action Effect on Probing Rate

10 20 30 40 50

10

0

20

30

40

50

Time (minutes)

Mean Probing Count

B

Control Treatment

Time (minutes)

20

0

40

60

80

100

120 Mean Landing Count

10 20 30 40 50

Control Treatment

A

Raid Dual Action Effect on Landing Rate

Figure 5. Time-series analysis of the Raid Dual Action effects 

on Ae. aegypti host-seeking activity: (A) Mean mosquito landing 

counts; (B) Mean number of mosquito probes; (C) Mean number 

of blood-fed mosquitoes.

Time (minutes)

Control Treatment

C

Mean Number of Blood-Fed Mosquitoes

Raid Shield Effect on Blood Feeding

3

5

1

0

2

4

6

10 20 30 40 50

5

0

10

15

20

25

30

35

40

Time (minutes)

Control Treatment

B

Mean Probe Counts

Raid Shield Effect on Probing Rate

10 20 30 40 50

Figure 6. Time-series analysis of the Raid Shield effects on 

Ae. aegypti host-seeking activity: (A) Mean mosquito landing 

counts; (B) Mean number of mosquito probes; (C) Mean number 

of blood-fed mosquitoes.

20

0

40

60

80

Time (minutes)

Control Treatment

A

Mean Landing Count

Raid Shield Effect on Landing Rate

10 20 30 40 50
January – June 2017 19

THE ARMY MEDICAL DEPARTMENT JOURNAL

the 50 minute test. Overall, knock-down effects for both 

products were minimal. Average knock-down was 1.08 

mosquitoes, with a maximum count of 4 (20%).

Semi-field (Enclosure)

Weather conditions at NECE, Naval Air Station, Jacksonville, Florida were hot and humid. Temperature and 

relative humidity recorded inside the semi-field enclosures averaged 25.5oC (max=29oC and min=21oC) 

and 86.8% (max=97% and min=66%). The prevailing wind direction was from the northeast with average daily wind speeds less than one kph. Regression 

analysis showed negative correlation of rep-averaged 

mosquito counts with temperature (C°) and humidity 

(%) ( r=-0.4637, df=16, P=.3406). Collections of mosquitoes from treatment (with a spatial repellent device) 

tents were significantly lower than the control (without a 

spatial repellent device) tents (P<.0001; Figure 7). The 

Raid Shield reduced mosquito entry into tents by 88%, 

while the Dual Action only decreased entry by 66%.

Comment

Various definitions and characterizations of spatial repellents have been reported in the literature. Dethier 33

defined spatial repellents as any stimulus constituting 

a vapor physical state which elicits an avoidance reaction. Gouck et al34 expanded the definition to include 

any compound or agent that can produce repellency at a 

distance. Nolen et al35 defined spatial repellents as “an 

inhibiting compound, dispensed into the atmosphere 

of three dimensional space which inhibits the ability of 

mosquitoes to locate and track a target such as a human 

or livestock.” Results from our investigation indicated 

that both of the SC Johnson spatial repellent products 

functioned as spatial repellents. The active ingredient used in both products was the synthetic pyrethroid 

transfluthrin. Unlike many of the other common pyrethroid chemicals, transfluthrin has a high vapor pressure which enables the chemical to passively vaporize 

at ambient temperature. Other pyrethroids such as permethrin require an external energy source for volatilization. In order to be a practical tool for Soldiers, a spatial 

repellent device must be effective, nontoxic, simple to 

construct and operate, and it should not require frequent 

maintenance. The 2 products used in this study are 

lightweight, inexpensive, contained low amounts of pesticide, and are easy to set up. From a logistics standpoint, 

these devices are available in high quantity and can be 

easily integrated into military logistic transportation 

platforms. For example, more than a hundred of these 

devices could be carried in a Soldier’s rucksack and the 

load effect would be minimal. The operation of these 

devices only consisted of placing it at the desired location and exposing the treated surface to the air (spray the 

product on the poster (Dual Action) or open the folded 

plastic (Raid Shield)). Compliance with mandated topical repellent use remains an issue in the military. Unlike 

topical repellents, these devices do not require reapplication by the user. Furthermore, the protection from the 

device benefits more than one person at a time.

Our laboratory (wind tunnel) evaluations of the SC Johnson spatial repellent products showed significant reductions in host-seeking behaviors (landing, probing and 

blood-feeding) of Ae. aegypti. However, reduction rates 

significantly decreased after 30 minutes. Previous studies using transfluthrin showed high efficacy with longer 

periods of effectiveness. Ogoma et al36 evaluated transfluthrin treated fabric (hessian cloth) against laboratoryreared An. arabiensis Patton. A single hessian strip (4.0 

m x 0.3 m) treated with 10 ml of transfluthrin reduced 

the mosquito attack rate on human volunteers by 99% 

and consistently conferred greater than 90% protective 

efficacy for a period of 6 months. Andrés et al37 evaluated transfluthrin dispensed from modified mosquito 

landing boxes against An. arabiensis in a semi-field system. The protective transfluthrin bubble provided 68.9% 

protection to human volunteers, and continued to show 

knockdown effects up to 3 weeks in postlaboratory testing. Govella et al38 tested transfluthrin treated strips in 

the field against An. gambiae. The strips conferred 99% 

protection and remained effective for over 3 months. One 

major limitation of our study was control of the amount 

of active ingredient used. Material Safety Data Sheets 

for both products showed that the amount of transfluthrin was less than 60 mg (less than 2% concentration). In 

comparison with previous studies, these amounts were 

relatively low. Additionally, it is important to note that 

the 2 products are marketed for indoor use.

Figure 7. Mean (±SE) mosquito capture rate per tent for control 

and treatments. Note: Bars tagged with the same letter are not 

significantly different.

20

0

40

60

80

100

120

Mean Number of Mosquitoes Collected

Raid Dual Action Raid Shield

a

a

b

b

Control Treatment
20 http://www.cs.amedd.army.mil/amedd_journal.aspx

Knockdown is the incapacitation of arthropods as a 

result of contact with a sublethal dose of pesticide. In 

this study, knockdown effects from transfluthrin exposure were low for both products. However, the relative 

knockdown of mosquitoes was consistently higher for 

the Dual Action device. This is particularly interesting 

due to the fact that the Dual Action device contained 

less transfluthrin. In the wind tunnel, mosquitoes often 

made direct contact with the Dual Action device. This 

was not observed with the Raid Shield device. The Dual 

Action product was designed with a multipurpose function of repelling insects and serving as a home freshener.

Our semi-field experiment demonstrated a tactical utility for spatial repellents. The BGS traps are commonly 

used by mosquito vector control districts to conduct 

surveillance and have been proven to be highly effective for collecting Ae. aegypti and Ae. albopictus. In 

this study, BGS traps baited with CO2

 were placed in 

military style tents and served as surrogates for humanbait collectors. Mosquito entry into military style tents 

was moderately reduced when using the spatial repellent devices. As expected due to environmental conditions, there was a significant variation in the amount 

of reduction. However, mosquito levels in treated tents 

remained well below that of control tents. This result 

suggests that the transfluthrin spatial repellents used in 

this study may have functioned as attraction inhibitors. 

A more likely explanation is that this behavior involved 

excito-repellency. For example, the mosquitoes could 

have detected and followed the CO2

 source inside the 

tent, but due to increased transfluthrin exposure, some 

of them quickly egressed from the tent to seek fresh air. 

Other studies suggest that placement of the device inside certain structures enhances the spatial repellent effect. Kawada et al39 evaluated metofluthrin-impregnated 

plastic strips against Ae. aegypti in houses located in 

My Tho City, Vietnam. Multiple regression analysis of 

environmental factors indicated that both an increase 

in average room temperature and a decrease in area of 

openings in rooms that were treated with metofluthrinimpregnated strips positively affected its spatial repellency. Transfluthrin exposure could have also negatively 

affected other key mosquito behaviors such as bloodfeeding and ovipositon. Unfortunately, the experimental design for the semi-field experiment did not include 

evaluations of these behaviors. Therefore, the effect of 

the transfluthrin on the mosquitoes that entered the experimental tents remains unknown.

This study suggests that spatial repellents have great potential for enhancing existing vector control and PPM 

efforts. However, sole reliance on this technology is 

not practical. Although spatial repellents have shown 

high efficacy rates in the field reaching levels of 99% 

protection, changing environmental conditions (wind, 

precipitation, temperature, and humidity) may present a major challenge for sustaining high efficacy rates. 

Traditional military approaches to mosquito control are 

based on the use of insecticides for area-wide abatement 

and PPM (topical repellents, treated uniforms, and bed 

nets). However, there is much concern among entomologists that these approaches may be severely restricted 

in the future due to insecticide resistance. Over the past 

few decades, there has been an overreliance on the use 

of pyrethroid insecticides for mosquito control. Global 

malaria eradication programs depend almost exclusively 

on pyrethroid-based indoor residual spraying and longlasting insecticidal nets (LLINs). Although this strategy 

has been viewed as having an important role in the recent reductions in global malaria mortality, major malaria vectors have developed resistance to this class of 

insecticides and the resistance alleles are very common 

in mosquito populations throughout the world. One way 

to overcome resistance is to rotate different classes of 

insecticides into a control program. Unfortunately, the 

acquisition of other classes of insecticides is often difficult and their cost is significantly higher than pyrethroids. Spatial repellents such as transfluthrin are readily available and are relatively inexpensive. A potential 

means of slowing the evolution of pyrethroid resistance 

may be the integration of spatial repellents into the program strategy.40 For example, the incorporation of a spatial repellent with LLINs would result in a 3-tier line of 

defense: (1) the spatial repellent, (2) contact pyrethroid, 

and (3) physical barrier (net). The spatial repellent would 

provide protection outside of the LLIN and contribute to 

reducing selection for resistance. Additional studies that 

explore the effects of integrating spatial repellents into 

DoD field vector control activities are warranted.

Acknowledgements

We thank SC Johnson and Company for their cooperation and assistance in providing the 2 products, Raid 

Dual Action Insect Repellent and Home Freshener and 

Raid Shield, used in the study.

We thank Anna Katrina Briley and Jason Fajardo for providing mosquitoes as well as for their assistance with the 

semi-field trials at NECE. We also thank Dr Muhammad 

Farooq for collecting the weather data at the Navy Entomology Center of Excellence.

Financial support for this project was provided by the 

Defense Health Program 6.7.

LABORATORY AND SEMI-FIELD EVALUATIONS OF TWO (TRANSFLUTHRIN) SPATIAL REPELLENT DEVICES

AGAINST AEDES AEGYPTI (L.) (DIPTERA: CULICIDAE)
January – June 2017 21

THE ARMY MEDICAL DEPARTMENT JOURNAL

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Authors

MAJ McPhatter is with the Entomology Branch, Walter Reed Army Institute of Research, Silver Spring, 

Maryland.

Dr Mischler is with the Entomology Branch, Walter Reed 

Army Institute of Research, Silver Spring, Maryland.

Dr Webb is with the Invasive Insect Biocontrol & Behavior Laboratory, US Department of Agriculture-Agricultural Research Service, Beltsville, Maryland.

Dr Chauhan is with the Invasive Insect Biocontrol & Behavior Laboratory, US Department of Agriculture-Agricultural Research Service, Beltsville, Maryland.

CPT Lindroth is with the Navy Entomology Center of 

Excellence, Jacksonville, Florida.

Dr Richardson is with the Navy Entomology Center of 

Excellence, Jacksonville, Florida.

Dr Debboun is the Director of the Mosquito & Vector 

Control Division, Harris County Public Health, Houston, 

Texas.

LABORATORY AND SEMI-FIELD EVALUATIONS OF TWO (TRANSFLUTHRIN) SPATIAL REPELLENT DEVICES

AGAINST AEDES AEGYPTI (L.) (DIPTERA: CULICIDAE)
January – June 2017 23

Zika virus (ZIKV), a mosquito-borne disease, was declared an international public health emergency by the 

World Health Organization (WHO) on February 1, 2016 

due to confirmed reports of various adverse health effects from multiple countries.1

 The state of Georgia 

Department of Public Health (DPH) lacked sufficient 

monitoring and surveillance capacity to properly assess 

the risk of ZIKV transmission, as well as trained staff to 

respond to potential local outbreaks of the disease. This 

limited capacity was a result of federal budget reductions 

that affected funding earmarked for mosquito monitoring and surveillance programs. Once a strong program 

that had a large number of county health departments 

conducting mosquito surveillance and several trained 

staff, the program had been reduced to one state level 

position and a few counties conducting surveillance. 

This case study describes and evaluates Georgia’s rapid 

response from the perspective of those challenges.

Overview

Zika virus is a mosquito-borne Flavivirus (family Flaviviridae) first discovered in 1947 in the Zika forest 

of Uganda from captive monkeys used in yellow-fever 

surveillance and later isolated in humans in 1952.2

 Unlike West Nile virus (WNV) that circulates between 

birds and mosquitoes with humans serving as dead-end 

hosts,3

 ZIKV can be transmitted from human-to-human 

via the bite of an infected Aedes spp mosquito.1

 From the 

1960s through the 1980s, the disease was found in mosquitoes from several Asian countries with few human 

cases. It eventually traveled east, resulting in a large outbreak on Yap Island in 2007 and additional outbreaks 

Georgia’s Collaborative Approach to

 Expanding Mosquito Surveillance in

 Response to Zika Virus: A Case Study

R. Christopher Rustin, DrPH, MT, REHS Sarbesh Pandeya, MPH

Deonte Martin, BS Haresh Rochani, DrPH, MPH, MBBS

Varadan Sevilimedu, MPH Rosmarie Kelly, PhD, MPH

Abstract

Zika virus (ZIKV) was declared an international public health emergency by the World Health Organization on 

February 1, 2016. Due to the known and estimated range of the ZIKV mosquito vectors, southern and central US 

states faced increased risk of ZIKV transmission. With the state of Georgia hosting the world’s busiest international 

airport, a climate that supports the ZIKV vectors, and limited surveillance (13 counties) and response capacity, the 

Department of Public Health (DPH) was challenged to respond and prevent ZIKV transmission. This case study 

describes and evaluates the state’s surveillance capacity before and after the declaration of ZIKV as a public health 

emergency.

Method: We analyzed surveillance data from the DPH to compare the geographical distribution of counties conducting surveillance, total number, and overall percentage of mosquito species trapped in 2015 to 2016. Counties 

conducting surveillance before and after the identification of the ZIKV risk were mapped using ArcMap 10.4.1. 

Using SAS (version 9.2) (SAS Institute, Inc, Cary, NC), we performed the independent 2 sample t test to test for 

differences in prevalence in both years, and a χ2 analysis to test for differences between numbers of species across 

the 13 counties. In addition, weighted frequency counts of mosquitoes were used to test (χ2 ) an association between 

major mosquito vector species and 7 urban counties. Lastly, using data from 2012-2016, a time-trend analysis was 

conducted to evaluate temporal trends in species prevalence.

Results: From 2015 to 2016, surveillance increased from 13 to 57 (338% increase) counties geographically dispersed 

across Georgia. A total of 76,052 mosquitoes were trapped and identified in 2015 compared to 144,731 (90.3% increase) in 2016. Significant differences between species (P<.001) and significant associations (P<.0001) between 7 

urban counties and major mosquito vectors were found. Significant differences in prevalence were found between 

several species and year highlighting species-year temporal trends.

Conclusions: The DPH collaborative response to ZIKV allowed a rapid increase in its surveillance footprint. Existing and new partnerships were developed with the military and local health departments to expand and share data. 

This additional surveillance data allowed DPH to make sound public health decisions regarding mosquito-borne 

disease risks and close gaps in data related to vector distribution.
24 http://www.cs.amedd.army.mil/amedd_journal.aspx

in French Polynesia in 2013.2 In 2015, Brazil reported 

unusual cases of rash and ultimately associated a high 

number of babies born with microcephaly and cases of 

guilliain-barré syndrome to ZIKV, resulting in the international public health emergency order.2

Research and analysis of previous outbreaks indicated 

that the primary vector responsible for transmitting 

ZIKV is the Aedes spp mosquito with the urban dwelling Ae. aegypti (L.) the likely vector in the Americas.4

 A 

daytime biter of humans, this mosquito prefers tropical 

to somewhat temperate climates and lays eggs in containers around urban areas.4

 A secondary ZIKV vector is Ae. albopictus (Skuse), the Asian tiger mosquito. 

This mosquito also lay eggs in containers, but feeds on 

humans and other animals, thus lowering the risk of 

transmission.4

 However, Ae. albopictus can survive in 

cooler areas, increasing its potential distribution across 

the United States. The Centers for Disease Control and 

Prevention (CDC), using several limited data sources, 

developed 2 maps (Figure 1) that estimate the potential 

geographic range of Ae. aegypti and Ae. albopictus.4

The potential distribution of these 2 vectors placed several states in the southern to central half of the United 

States on alert for potential transmission of ZIKV and 

raised the alarm for public health officials in Georgia. 

Unfortunately, the maps demonstrate the lack of surveillance across the country, as it only estimates the 

potential range of the 2 ZIKV vectors. This data gap is 

a result of public health budget cuts and lack of priority 

placed on mosquito surveillance in the last decade. Consequently, this negatively affected the state’s ability to 

critically assess the actual risks of ZIKV transmission 

for its citizens.

National Funding Trends

Understanding national funding trends is important to 

explaining the gaps in surveillance across the United 

States. The primary mission of public health and mosquito control programs is to inform, prevent, and protect the public from injury and disease. This mission is 

achieved through disease and vector surveillance programs that provide critical data used to quickly respond 

and control threats. Mosquito surveillance, coupled 

with clinical (human and animal) surveillance programs, are critically important because they can detect 

the abundance and distribution of vectors, monitor for 

viral diseases, aid in quantifying human risks, and predict changes in the dynamics of disease transmission.6-8

Unfortunately, funding and support of surveillance programs have decreased over time, leaving a patchwork 

of jurisdictions conducting surveillance and resulting in 

significant gaps in vector data.9

West Nile virus (WNV), the last major new and exotic arboviral outbreak in the United States, generated significant media attention and heightened the public’s fear.10-12

This outbreak highlighted a general lack of capacity for 

public health and mosquito control agencies to conduct 

human and vector surveillance and rapidly respond to 

disease events.13 To improve detection, monitoring, and 

control capacity for WNV (which eventually spread to 

48 states), the CDC provided significant funding to all 

50 states and 6 major cities through its Epidemiology 

and Laboratory Capacity (ELC) grant.13 This funding 

was eventually expanded to cover over 20 mosquitoborne and tick-borne diseases and allowed states and 

jurisdictions to increase human and vector surveillance, 

mosquito trapping and identification, viral testing, avian 

surveillance systems; and develop response plans.14

In 2002, approximately $34.7 million was provided to 

states to fund these important programs, but by 2014, 

this had been reduced by approximately 75% to $9.2 

GEORGIA’S COLLABORATIVE APPROACH TO EXPANDING MOSQUITO SURVEILLANCE

IN RESPONSE TO ZIKA VIRUS: A CASE STUDY

Figure 1. Estimated range of Aedes albopictus and Aedes aegypti in the United States, 2016. The maps represent the best 

estimate by CDC of the potential range of the named mosquitoes. Maps courtesy of the Centers for Disease Control and 

Prevention.4
January – June 2017 25

THE ARMY MEDICAL DEPARTMENT JOURNAL

million.14 A CDC report based on a follow-up survey 

conducted by the Council of State and Territorial Epidemiologists indicated that since 2005, 22% of funded 

jurisdictions eliminated active human surveillance, 13% 

eliminated mosquito surveillance, 70% eliminated mosquito trapping, and 64% stopped avian surveillance.13

Since 2005, the number of counties conducting routine 

mosquito surveillance in Georgia was reduced by 78%, 

from 60 to 13 counties, and all public health funding for 

arboviral testing and avian surveillance was eliminated 

(R. Kelly, unpublished data*). This report concluded 

that the state’s ability to rapidly detect and respond to 

emerging disease threats is “compromised,”13 thus placing states, including Georgia, at a disadvantage in responding to vector-borne diseases.

Georgia Preparedness

Georgia, like many states, faces several challenges to 

monitoring and responding to emerging arboviral diseases, with ZIKV highlighting this issue. In addition 

to the CDC estimating that the range of ZIKV vectors 

spans the entire state, several additional issues raise 

the risk of ZIKV transmission. First, Georgia is home 

to the world’s busiest international airport, thus hosting 

visitors and tourists from ZIKV endemic countries. Additionally, many Georgia citizens travel to tourist destinations that have active ZIKV circulating. This raised 

the risk that a traveler could become infected and return 

to Georgia, thus spreading ZIKV to local mosquitoes. 

Second, Georgia lacks statewide surveillance and vector control which are 

limited to just 13 counties conducting 

surveillance and 6 counties providing 

comprehensive mosquito control services. Third, previous surveillance has 

determined that Ae. albopictus is present in every county,*15 but with limited 

surveillance, the true distribution of Ae. 

aegypti is unknown having only been 

recently found in 2 counties. Ae. aegypti had been a common species in 

Georgia until the introduction of Ae. albopictus in the 1990s. This lack of surveillance data prevents accurate quantification of human risks. Lastly, due to 

the federal funding cuts, the DPH has 

limited capacity to conduct comprehensive surveillance and mapping of vector 

abundance and evaluate insecticide resistance, and limited ability to provide 

emergency vector control.

These challenges are compounded by the fact that Georgia’s population is over 10 million people and growing 

(Figure 2), and has the largest number of counties (159), 

second only to the state of Texas.16 These counties range 

from urban to suburban, with approximately 108 (68%) 

of the counties classified as rural (less than 35,000 population).17 With the population of the state steadily increasing, and a public health program with limited resources, 

more people are potentially at risk of exposure to emerging vector-borne diseases. However, faced with these 

challenges, the DPH used the expertise of its State Entomologist and the environmental health program to evaluate its weaknesses and implement a rapid, collaborative 

response to the threat of ZIKV.

Purpose

The purpose of this case study was to describe and compare the DPH’s mosquito surveillance capacity in 2015 

before ZIKV was declared a public health emergency to 

the 2016 surveillance capacity following that declaration. 

Statistical comparisons were made between years on the 

number of counties conducting surveillance and differences in prevalence and species, in addition to a time 

trend analysis of mosquito species distribution. These 

data were used to evaluate the state’s rapid response to 

the threat of ZIKV and the risk of autochthonous vector 

transmission based on the presence of the ZIKV vectors.

Methods

Mosquito surveillance trapping data provided by the 

DPH and surveillance data collected in collaboration 

Figure 2. Population of the state of Georgia by year, 2005-2015. Georgia Population (millions)

2005 2006 2007 2008 2009 2010 2011 2012 2013 2014 2015

10.0

10.5

9.5

9.0

8.5

8.0

Projected: 10.2 Million

*Unpublished arboviral summary statistics report prepared quarterly and annually by author R. Kelly and distributed internally 

within DPH, public health departments, MC agencies, and academic partners.
26 http://www.cs.amedd.army.mil/amedd_journal.aspx

with DPH were analyzed. The geographical distribution 

of counties conducting surveillance, total number, and 

overall percentage of mosquito species collected in 2015 

were compared to 2016 data. The distribution of counties conducting surveillance was mapped using ArcMap 

10.4.1 (Esri, Inc, Redlands, CA).

Statistical comparisons were made for the 13 counties 

conducting mosquito surveillance between 2015 and 

2016 to test for differences in response before and after the ZIKV emergency declaration. To test differences 

in prevalence (number of mosquitoes trapped/100,000 

population) in both years, the overall number of mosquitoes was compared using the independent 2 sample 

t test. A χ2 analysis was performed to test for statistical 

differences between numbers of species trapped across 

all 13 counties. In addition, weighted frequency counts 

of mosquitoes were used to conduct a test of association 

(χ2 ) between the major vector species of mosquitoes and 

the geographic county in 7 urban counties. The counties included in this χ2 analysis were Chatham, Fulton, 

Glynn, Liberty, Lowndes, Muscogee, and Richmond. 

Lastly, a time-trend analysis was conducted using surveillance data from 2012-2016 to test differences in species prevalence over time. Statistical significance was 

established at the significance level 0.05.

Results

In 2015, prior to the declaration that ZIKV represented a 

public health emergency, Georgia had 13 counties conducting surveillance, with the DPH medical entomologist 

(one full time equivalent/statewide) providing routine 

surveillance in 4 of those counties. In 2016 (March-December), the DPH expanded surveillance to 57 counties 

(338% increase) geographically dispersed in urban and 

rural areas (Figure 3).

This rapid expansion of surveillance was a result of hiring new staff and collaborating with the local health departments (LHDs) and the military. Table 1 shows that 

76,052 mosquitoes were trapped and identified in 2015 

compared to 144,731 mosquitoes trapped in 2016, representing a 90% increase. Forty-four mosquito species 

were identified in both years with Culex quinquefasciatus (Say), Georgia’s primary WNV vector, representing 

the highest percentage of mosquitoes trapped in both 

years (79.45% and 62.53% respectively). In reference to 

ZIKV vectors, Ae. aegypti represented only 0.108% and 

0.018% respectively of the total mosquitoes trapped each 

year, and were found in Muscogee County only, home to 

Fort Benning. Aedes albopictus represented only 1.50% 

and 3.703% of the total mosquitoes trapped respectively 

each year, reported from 11/13 (84%) counties in 2015 

and 42/57 (74%) counties in 2016.

Overall Mosquito Prevalence

To test differences in overall prevalence (number of 

mosquitoes/100,000 population) of mosquitoes trapped 

in both years from the 13 counties that have historically 

conducted surveillance, the number of mosquitoes was 

compared using the independent 2 sample t test. The 

value of the number of mosquitoes was normalized 

GEORGIA’S COLLABORATIVE APPROACH TO EXPANDING MOSQUITO SURVEILLANCE

IN RESPONSE TO ZIKA VIRUS: A CASE STUDY

Figure 3. Georgia counties conducting mosquito surveillance in collaboration with the Georgia Department of Public Health in 

2015 (13 counties) and 2016 (57 counties).

 2015 Surveillance

 No Surveillance No Surveillance

 2016 Surveillance

No Surveillance

2015 2016
January – June 2017 27

THE ARMY MEDICAL DEPARTMENT JOURNAL

using the log transformation and the normality 

was confirmed with Kolmogorov-Smirnoff test. 

While the total number of mosquitoes trapped increased from 2015 to 2016 for the 13 counties, this 

analysis shows there was no statistically significant 

differences (P=.7901) in the overall prevalence of 

mosquitoes trapped before and after ZIKV was 

declared an international public health emergency 

(Table 2A).

Table 2B shows the significance of the t test assuming equal and unequal variances. Table 2C shows 

the results for the test of equality of variances between the 2 samples

Species Analysis

To test for differences in the prevalence of individual mosquito species trapped in 2015 versus 2016 

for the 13 counties, a χ2 analysis was performed. 

Results show significant differences (P<.001) in 

the distribution of all mosquito species trapped in 

those years (Table 1). However, while there were 

significant differences in prevalence, this does not 

tell us the behaviors associated with individual 

species.

By combining both years (2015, 2016) of mosquito 

data, a secondary χ2 analysis was conducted to 

test for an association between Georgia’s primary 

arboviral vector species and urban counties with 

a large population of people at risk. Adjusting for 

inconsistent or missing data, weighted frequency 

counts of mosquitoes were used in the analysis. 

While the previous t test results (Table 2) indicate 

that overall difference in prevalence of mosquitoes 

is not significant between 2015 and 2016, there 

Table 2A. The 2-sample t test for difference in 

mosquito prevalence between 2015 and 2016.

Year Method Mean 95% CI

2015 6.34 4.79, 7.88

2016 6.63 4.86, 8.39

Diff (1-2) Pooled -0.29 -2.51, 1.93

Diff (1-2) Satterthwaite -0.29 -2.51, 1.94

Table 2B. The significance of the t test assuming 

equal and unequal variances.

Method Variances df t value P value

Pooled Equal 24 -0.27 .7901

Satterthwaite Unequal 23.589 -0.27 .7902

Table 2C. Results for the test of equality of variances between the 2 samples.

Method Num df Den df F-value P value

Folded F 12 12 1.30 .65

Table 1. Mosquito Species Collected, 2015-2016.

Species* 2015

Totals

13 Counties

Percentages

2016

Totals

57 Counties

Percentages

Ae. aegypti 82 0.108% 26 0.018%

Ae. albopictus 1,141 1.500% 5,360 3.703%

Ae. cinereus 0 0.000% 4 0.003%

Ae. japonicus 0 0.000% 1 0.001%

Ae. vexans 162 0.213% 6,536 4.516%

Aedes/Ochlerotatus spp. 6 0.008% 120 0.083%

An. barberi 0 0.000% 1 0.001%

An. crucians 25 0.033% 1,873 1.294%

An. punctipennis 26 0.034% 486 0.336%

An. quadrimaculatus 61 0.080% 265 0.183%

Anopheles spp. 5 0.007% 134 0.093%

Cq. perturbans 1,265 1.663% 5,969 4.124%

Cs. inornata 130 0.171% 14 0.010%

Cs. melanura 906 1.191% 996 0.688%

Culex spp. 4,996 6.569% 10,830 7.483%

Cx. coronator 262 0.345% 604 0.417%

Cx. erraticus 300 0.394% 2425 1.676%

Cx. nigripalpus 5,657 7.438% 11,071 7.649%

Cx. peccator 0 0.000% 12 0.008%

Cx. quinquefasciatus 60,423 79.450% 90,505 62.533%

Cx. restuans 100 0.131% 389 0.269%

Cx. salinarius 350 0.460% 2,746 1.897%

Cx. territans 1 0.001% 33 0.023%

Ma. titillans 0 0.000% 98 0.068%

Oc. japonicus 8 0.011% 52 0.036%

Oc. atlanticus 1 0.001% 757 0.523%

Oc. canadensis 0 0.000% 117 0.081%

Oc. fulvus pallens 0 0.000% 1 0.001%

Oc. infirmatus 2 0.003% 45 0.031%

Oc. mitchellae 0 0.000% 9 0.006%

Oc. sticticus 0 0.000% 31 0.021%

Oc.taeniorhynchus 0 0.000% 5 0.003%

Oc. triseriatus 25 0.033% 78 0.054%

Or. signifera 3 0.004% 23 0.016%

Ps. ciliata 0 0.000% 25 0.017%

Ps. cyanscens 2 0.003% 30 0.021%

Ps. columbiae 88 0.116% 332 0.229%

Ps. ferox 10 0.013% 106 0.073%

Ps. howardii 3 0.004% 34 0.023%

Psorophora spp. 6 0.008% 0 0.000%

Tx. rutilus 1 0.001% 52 0.036%

Ur. iowii 0 0.000% 13 0.009%

Ur.sapphrina 2 0.003% 115 0.079%

Unknown 3 0.004% 2,408 1.664%

Total 76,052 144,731

*Data for distribution of all species statistically significant (P≤.05).
28 http://www.cs.amedd.army.mil/amedd_journal.aspx

is a significant association (P<.0001) between urban 

counties and major vector species as shown in Table 3. 

However, while χ2 tests can provide information on distribution of species between years, it does not provide 

detailed information on behavior of individual species, 

trends, and patterns over time.

Time Trend Species Analysis

A time-trend species analysis using generalized linear models (Proc GLM in SAS 9.2) was conducted on 

mosquito data from 2012 to 2016 to evaluate temporal 

trends in prevalence of mosquitoes. Statistically significant differences were observed over the 5-year period 

in species prevalence (P<.0001). While time (in years) 

was not significantly associated with change in overall 

prevalence, (P<.1219) its differential effect on species 

specific change in prevalence was significant (interaction term – P<.0001) (Table 4).

Looking more closely at the specific species estimates in 

Table 5, it can be seen that certain species had significantly different prevalence compared to the reference group, 

ie, Oc. sticticus (Meigen), throughout the 5-year period. 

This reference group was chosen due to low numbers 

of species trapped. These species were Ae. albopictus

(S.) (P<.0001), Ae. vexans (Meigen) (P=.0240), Cq. perturbans (Walker) (P<.0001), Cs. melanura (Coquillett) 

(P<.0001), Culex spp. (P<.0001), Cx. coronator (Dyer 

and Knab) (P=.0463), Cx. erraticus (Dyer and Knab) 

(P<.0001), Cx. nigripalpus (Theobald) (P<.0001), Cx. 

quinquefasciatus (Say) (P<.0001), Cx. restuans (Theobald) (P=.0004), Cx. salinarius (Coquillett) (P<.0001) 

and Oc. triseriatus (Say) (P<.0001).

For associated species-year trends in Table 5, Cs. melanura (P<.0042), Culex spp. (P<.0013), Cx. erraticus

(P <.0271), Cx. nigripalpus (P<.0074), Cx. restuans

(P<.0433) and Cx. salinarius (P<.0266) were found to 

have a significant difference in change of prevalence with 

each year compared to the reference species Oc. sticticus.

Comment

The public health entomology program, which includes 

mosquito surveillance, falls under the DPH Environmental Health (EH) Section. This program is managed 

by a medical entomologist who holds graduate degrees 

in public health and entomology and who was originally hired under Epidemiology and Laboratory Capacity 

funding in 2002 to manage the WNV outbreak. The 

state medical entomologist provides technical assistance 

and consultation to LHDs, mosquito control agencies 

and the general public and was largely responsible for 

overseeing Georgia’s rapid response to ZIKV. In 2015, 

surveillance and identification of mosquitoes was limited to just 13 counties in Georgia. With this limited 

data, DPH provided quarterly surveillance reports 

to partner agencies for planning and risk assessment 

purposes.

GEORGIA’S COLLABORATIVE APPROACH TO EXPANDING MOSQUITO SURVEILLANCE

IN RESPONSE TO ZIKA VIRUS: A CASE STUDY

Table 3A. Weighted frequency counts and corresponding weighted row percentages of important mosquito species stratified by county.

Chatham Fulton Glynn Liberty Lowndes Muscogee Richmond Total

Ae. aegypti 2

1.67%

2

1.67%

2

1.67%

2

1.67%

2

1.67%

108

90.00%

2

1.67%

120

Ae. albopictus 26

0.70%

871

23.46%

72

1.94%

53

1.43%

177

4.77%

653

17.59%

1,861

50.12%

3,713

Ae. vexans 2

0.07%

94

3.23%

2

0.07%

2

0.07%

2

0.07%

101

3.47%

2,704

93.02%

2,907

Cs. melanura 196

10.36%

2

0.11%

2

0.11%

2

0.11%

1,686

89.11%

2

0.11%

2

0.11%

1,892

Culex spp. 13,985

99.30%

28

0.20%

2

0.01%

2

0.01%

2

0.01%

17

0.12%

48

0.34%

14,084

Cx. nigripalpus 1,417

8.56%

2

0.01%

529

3.20%

2

0.01%

13,556

81.90%

2

0.01%

1,044

6.31%

16,552

Cx. quinquefasciatus 58,184

42.09%

8,472

6.13%

41,910

30.32%

85

0.06%

23,776

17.20%

778

0.56%

5,031

3.64%

138,236

Total 73,812 9,471 42,519 148 39,201 1,661 10,692 177,504

Table 3B. Chi-square value with corresponding P value (α=0.05). Statistic df Value P value

χ2 36 143,393 <.0001

Table 4. Time trend analysis showing the type III sums of squares 

and corresponding P values (α=0.05) for the variables species, 

year, and the interaction term.

df Type III SS Mean Square F-value P value

Species 41 1589.943667 38.779114 14.25 <.0001

Year 1 6.521391 6.521391 2.40 .1219

Year species 41 318.828534 7.776306 2.86 <.0001
January – June 2017 29

THE ARMY MEDICAL DEPARTMENT JOURNAL

When ZIKV was declared an international public 

health emergency,1

 the DPH recognized its weaknesses and formed a core team of professionals to 

develop and lead a response. This diverse team was 

comprised of EH, medical entomology, medical epidemiology, communications, emergency preparedness, and leadership. An assessment was conducted to 

ascertain agency strengths, weaknesses, and overall 

needs to develop a response. This evaluation assessed 

geographical gaps in mosquito data and surveillance 

staff needs, opportunities for new or expanded partnerships that could enhance a response and facilitate 

data sharing, training of existing staff, and funding 

needs. Leadership made the ZIKV response a priority 

and identified existing grant funding, which significantly contributed to a rapid response. The assessment 

guided DPH in utilizing those funds to hire additional 

surveillance staff and assign them regionally across 

the state, updated existing surveillance and response 

plans originally written for WNV, provided rapid 

training to new and existing staff across the state in 

surveillance and response, purchased equipment, and 

expanded collaborations with the military to share 

data and respond to this threat.

As the results in Table 1 indicate, DPH successfully 

and rapidly expanded its surveillance footprint by 

338%, from 13 counties in 2015 to 57 counties in 2016, 

representing 36% of the state. While the overall number 

of counties conducting surveillance seems low, given 

there are 159 counties, it should be noted that this rapid 

expansion occurred in just 8 months (May-December) 

and all major urban population centers have active surveillance. In addition, the distribution of surveillance 

covered all regions of the state. This expansion led to 

the overall number of mosquitoes trapped increasing 

by 90.3%, from 2015 to 2016, allowing better decision 

making. Rapid expansion was achieved because the 

DPH leadership made the ZIKV response a top priority and streamlined the hiring and purchasing processes. 

Prior to hiring new staff (March-April), the environmental health program updated its WNV surveillance 

and response plan by tailoring it for ZIKV. In addition, 

a training curriculum, standard operating procedures, 

job description for surveillance staff, and regions were 

established. The DPH collaborated with regional public 

health departments to provide office and storage space 

for the new surveillance staff, a critical component for a 

successful program.

In April and May, 5 new surveillance staff members 

were hired and provided 2 weeks of just-in-time training on mosquito surveillance techniques and identification, with priority placed on ZIKV vectors, risk 

communication, data management, and vector control. 

Each new staff member was assigned a region to cover 

and provided with surveillance and response equipment 

(traps, larvicides, backpack sprayers, microscope, educational material). In addition, the state entomologist provided ongoing training and consultation throughout the 

year, and assisted new staff with establishing surveillance 

sites throughout their region. To assist the new regional 

staff, the DPH provided training, surveillance equipment, 

and funding to existing LHD environmental health staff 

across the state, efforts which were invaluable in expanding the surveillance footprint across Georgia.

Historically, Georgia’s mosquito surveillance objectives 

were driven by the threat of WNV, and mosquito trapping was prioritized to capture the primary WNV vector, Cx. quinquefasciatus. This is highlighted in Table 1, 

with Cx. quinquefasciatus as the highest percentage of 

mosquitoes trapped overall in 2015 and 2016 at 79.4% 

(60,423) and 62.5% (90,505) respectively. However, 

the threat of ZIKV required surveillance staff to shift 

some focus away from the WNV vector and prioritize 

trapping in urban/suburban areas to target containerbreeding Ae. aegpyti and Ae. albopictus. While overall 

counts and percentages of both ZIKV vectors were low 

in 2015 and 2016, it is important to note that this shift 

in surveillance focus is demonstrated in Table 1 by an 

Table 5. The results of the PROC GLM (SAS 9.2) procedure showing 

changes in the log of mosquito prevalence by year and the 

differential trends in log of mosquito prevalence by year.*

Parameter Estimate Standard 

Error

t value P value

Intercept 0.18 0.61 0.29 .76

Ae. albopictus 5.34 1.20 4.44 <.0001

Ae. vexans 1.84 0.81 2.26 .02

Cq. perturbans 2.84 0.71 4.00 <.0001

Cs. melanura 3.03 0.72 4.21 <.0001

Culex spp. 4.02 0.75 5.33 <.0001

Cx. coronator 1.50 0.75 1.99 .0463

Cx. erraticus 4.00 0.79 5.01 <.0001

Cx. nigripalpus 4.07 0.68 5.91 <.0001

Cx. quinquefasciatus 6.79 0.66 10.16 <.0001

Cx. restuans 2.48 0.69 3.55 .0004

Cx. salinarius 2.97 0.69 4.30 <.0001

Oc. triseriatus 1.62 0.70 2.29 .02

Oc. sticticus Ref

year species Cs. melanura -1.29 0.45 -2.87 .004

year species Culex spp. -1.51 0.46 -3.23 .001

year species Cx. erraticus -1.10 0.50 -2.21 .02

year species Cx. nigripalpus -1.15 0.43 -2.68 .007

year species Cx. restuans -0.88 0.43 -2.02 .04

year species Cx. salinarius -0.95 0.43 -2.22 .02

year species Oc. sticticus Ref

*Only results significant at α=0.05 are shown.
30 http://www.cs.amedd.army.mil/amedd_journal.aspx

increase in Ae. albopictus trapped (1.5% to 3.7%) and 

a decrease in WNV vectors trapped (79.4% to 62.5%). 

Aedes aegypti, the primary ZIKV vector, was trapped in 

one county (Muscogee) for both years and overall numbers decreased from 82 mosquitoes in 2015 to 26 in 2016, 

representing an 83% decrease. While surveillance has 

been limited, Ae. aegpyti was only found in Muscogee 

County and Chatham County in the last decade, suggesting that Ae. albopictus has outcompeted this species 

given its propensity for surviving in surburban/urban 

areas.18

The next 4 highest percentages of mosquitoes trapped in 

2016 were Cx. nigripalpus, Culex spp, Ae. vexans, and 

Cq. perturbans. The number of Ae. vexans exploded in 

2016 to 6,583 mosquitoes compared to 2015 at 162. This 

increase came in the aftermath of Hurricane Matthew, 

which left behind ideal breeding conditions for this inland floodwater species, resulting in standing water in 

low-land and grassy areas of southeast Georgia. It’s important to point out that the top 5 mosquitoes captured 

are major vectors of Eastern Equine Encephalitis (EEE), 

St. Louis Encephalitis (SLE), and WNV, either serving 

to amplifying the virus enzootically, or as a primary or 

bridge vector to humans.19-21

Statistical Analysis

To determine if there were any statistical differences or 

associations in the DPH surveillance program before 

and after the ZIKV emergency declaration, a series of 

statistical tests were run comparing data between the 

same 13 counties historically conducting surveillance. 

The t test examined the overall prevalence of mosquitoes per 100,000 population. While this test did not 

show statistically significant differences in prevalence, 

it should be noted that the overall number of mosquitoes 

trapped increased by 66%, from 76,052 to 126,584 mosquitoes in these 13 counties. This increase demonstrated 

the rapid expansion of surveillance in response to ZIKV.

A series of χ2 analyses were conducted to test for any 

differences between both years for the same counties at 

the species level. Overall, there were significant differences found in several of the mosquito species trapped 

between years. Of importance to ZIKV, there was a 

statistically significant increase in Ae. albopictus from 

2015 to 2016 and a statistically significant decrease in 

Ae. aegypti between those years. In addition, there was 

a statistically significant decrease in the number of Cx. 

quinquefasciatus. This was likely due to a shift in focus on targeting the ZIKV vectors which have different 

breeding habitats. However, Georgia experienced a dry 

late spring and summer with abnormally cooler nights, 

which could have affected both vector species.

When Georgia’s primary disease vector species was statistically compared to large urban counties, a significant 

association between species and county was found. This 

is critically important because the majority of the state’s 

population is found in these large urban counties, putting more people at risk for arboviral diseases and demonstrating the ongoing need for surveillance.

To determine if there were any temporal trends in the 

distribution of species over time, a time-trend analysis 

was conducted on mosquito data from 2012 through 

2016. This analysis demonstrated that over a 5-year period, there were significant differences in prevalence for 

several important disease vector species in Georgia that 

transmit WNV (Cx. quinquefasciatus) , EEE (Cs. melanura, Ae. vexans, Culex spp), SLE (Cx. nigripalpus)

and LaCrosse encephalitis (Oc. triseriatus), in addition 

to nuisance species. Of note is the significant prevalence 

of Oc. triseriatus over the 5-year period. This mosquito represents Georgia’s primary vector for LaCrosse 

encephalitis and demonstrates potential risk, as it has 

been trapped consistently over the 5-year period. It can 

adapt to its surroundings and lay eggs in forest or urban 

container environments,22 highlighting the critical importance of continued surveillance for this species and 

other major disease vectors. Knowing these temporal 

trends allows public health and mosquito control professionals to predict risks and better prepare.

Military Collaboration

While Georgia was able to expand its surveillance capacity rapidly with internal resources, it is important 

to point out the vital collaborations and contributions 

made by the military. These collaborations were essential because military personnel travel to ZIKV endemic 

countries and many of them live off base, necessitating 

a need to partner and share data. In past years when 

WNV was the focus of concern, the DPH Vector-borne 

and Zoonotic Section formed a cooperative agreement 

with the US Army Center for Health Promotion and 

Preventive Medicine (USACHPPM) to share mosquito 

surveillance and testing data collected within the State. 

Unfortunately, when USACHPPM-South moved its 

headquarters from Georgia to Fort Sam Houston, Texas, 

that collaboration was largely lost.

In 2016, as mosquito surveillance was being increased 

in Georgia, a connection with the military was sought 

again. Not having one central agency made this process 

somewhat difficult, as each base had to be approached 

individually, but ultimately several connections were 

made. As one example, Dobbins Air Reserve Base 

Bioenvironmental Engineers (BEE) partnered with the 

DPH as part of the Department of Defense initiative to 

GEORGIA’S COLLABORATIVE APPROACH TO EXPANDING MOSQUITO SURVEILLANCE

IN RESPONSE TO ZIKA VIRUS: A CASE STUDY
January – June 2017 31

THE ARMY MEDICAL DEPARTMENT JOURNAL

combat ZIKV and other mosquito-borne diseases. One 

of DPH’s new vector surveillance coordinators, along 

with BEE personnel, laid traps at the Dobbins firing 

range, an area of frequent mosquito complaints. Mosquitoes collected were identified, and education, source 

reduction, and chemical control were used to reduce 

mosquito numbers in the area and reduce risk to military personnel. Follow up surveillance was done, and 

plans were made to continue this partnership in 2017. 

This important partnership was published by the US Air 

Force News Service as shown in Figure 4.23

In addition, Fort Benning, Fort Stewart, Hunter Army 

Airfield, and Robins Air Force Base agreed to share mosquito surveillance and testing data with the DPH. This 

not only provides additional information to the DPH concerning disease risks, but also provides mosquito control both on and off the bases with the information they 

need to reduce mosquito populations and disease risk. It 

should be noted that Fort Gordon has had a long-standing 

collaboration with the Richmond County Mosquito Control Program, a division of the local health department.

Conclusion

The collaborative response to ZIKV allowed DPH to rapidly increase its surveillance footprint across the state 

and train new and existing staff on outbreak response. 

With these new monitoring and response capabilities, 

the DPH can make sound public health decisions regarding disease risks and quickly respond to local outbreaks 

of ZIKV or other vector borne diseases.

Enhanced Capabilities

The Georgia DPH was able to rapidly expand its surveillance capacity statewide because its leadership recognized the seriousness and potential impact of ZIKV on 

the state and prioritized vector-borne diseases and surveillance. This allowed the agency to maximize existing 

resources to expand surveillance capacity and reignite 

historic and develop new collaborations with 

various entities, most importantly the LHDs 

and the military. This expanded surveillance 

network provided a clearer picture of the types 

of mosquitoes potentially exposing the public 

to mosquito-borne diseases and allowed DPH 

to better quantify risks and provide public education. Statistical analysis of the data validates 

the need for ongoing expanded surveillance.

In evaluating the risk of ZIKV transmission, 

recent historic data for the primary vector of 

ZIKV, Ae. aegypti, was isolated to just 2 counties in Georgia. Expanded surveillance in 2016 

confirmed a low presence of Ae. aegypti having 

been found in just one county, suggesting the 

primary vector for Zika has been displaced by 

Ae. albopictus. This also suggests a reduced risk 

of autochthonous transmission of Zika virus in 

Georgia due to the affinity of Ae. Albopictus for 

feeding on both humans and animals. However, 

this should be interpreted with caution due to 

unstandardized reporting techniques for each 

county, lack of systematic surveillance in every county, and a dry spring and summer that 

reduced overall number of mosquitoes trapped. 

The DPH is working with all counties to improve the quality of data reported, and will continue to 

assess the abundance and distribution of Ae. aegypti

each year to evaluate risks. In addition, while DPH expanded surveillance to 57 counties which comprise approximately 36% of the state’s 159 counties, the agency 

continues to work with the LHDs to establish additional 

surveillance sites with the goal of 100% of counties conducting mosquito surveillance. This will allow better 

interpretation of overall mosquito abundance and distribution across the state.

Public Health Implications

By increasing the number of counties involved in surveillance and knowing the specific species of mosquitoes throughout the state, the DPH can better predict 

and quantify the risks of disease transmission in specific 

Figure 4. Georgia Department of Public Health vector surveillance coordinator installing mosquito traps at the Dobbins Air Reserve Base.23
32 http://www.cs.amedd.army.mil/amedd_journal.aspx

regions of Georgia. However, increasing and maintaining surveillance comes with its own challenges, with 

stable funding being the largest obstacle to a robust 

surveillance system. The DPH was able to prioritize 

existing funding to kick start a rapid response and was 

subsequently awarded grants from the CDC specifically 

for ZIKV, but these funding streams are temporary and 

allocated for a specific disease. It is critically important 

that funding for monitoring, surveillance, and response 

becomes a permanent federal funding source, and that it 

does not follow the same path as WNV funding reductions of the past. If so, the United States may not be prepared to respond to the next emerging arboviral disease, 

and the results could be disastrous for a population that 

has no natural immunity.

References

1. World Health Organization. Zika virus and complications [internet]. Updated February 2, 2017. 

Available at: http://www.who.int/emergencies/zikavirus/en/. Accessed February 2, 2017.

2. World Health Organization. Zika virus [internet]. 

Updated September 6, 2016. Available at: http://

www.who.int/mediacentre/factsheets/zika/en/. Accessed February 2, 2017.

3. Randle YH, Freeman CB, Jackson M, Reyna M, 

Debboun M. 2014: A record-breaking year for West 

Nile virus positive mosquito pools in Harris County and the City of Houston, Texas. US Army Med

Dep J. October-December 2016;1-8. Available at: 

http://www.cs.amedd.army.mil/FileDownloadpublic.aspx?docid=ede3a0ed-f2ff-4ed1-9414-7e04d

b25e724. Accessed March 16, 2017.

4. Centers for Disease Control and Prevention. Zika 

Virus: Potential range in the United States [internet]. Updated January 19, 2017. Available at: https://

www.cdc.gov/zika/vector/range.html. Accessed 

February 2, 2017.

5. Moore CG, Mitchell CJ. Aedes albopictus in the 

United States: 10-year presence and public health 

implications. Emerg Inf Dis. 1997;3(3):329-334.

6. Vasquez-Prokopec GM, Chaves LF, Ritchie 

SA, Davis J, Kitron U. Unforeseen costs of cutting surveillance budgets. PLoS Negl Trop Dis. 

2010;4(10):e858. doi: 10.1371/journal.pntd.0000858.

7. Flores C. Mosquito surveillance for effective mosquito population control [internet]. Vector Disease 

Control International. Available at: http://www.

vdci.net/blog/mosquito-surveillance-for-effectivemosquito-population-control. August 19, 2015. Accessed February 2, 2017.

8. Moore CG, McLean RG, Mitchell CJ, et al. Guidelines for arbovirus surveillance programs in the 

United States [internet]. Centers for Disease Control and Prevention. April 1993. Available at: https://

www.cdc.gov/ncezid/dvbd/pdf/arboguid_508.pdf. 

Accessed February 2, 2017.

9. Council of State and Territorial Epidemiologists. 

Assessment of capacity in 2012 for the surveillance, 

prevention, and control of West Nile virus and 

other mosquito-borne virus infections in state and 

large city/ county health departments and how it 

compares to 2004 [internet]. February 2014. Available at: http://www.cste2.org/docs/VBR.pdf. Accessed February 2, 2017.

10. Asnis DS, Conetta R., Teixeira AA, Waldman G, 

Sampson BA, The West Nile virus outbreak of 

1999 in New York: the Flushing hospital experience. Clin Infect Dis. 2000;30:413-418.

11. Asnis DS, Conetta R, Waldman G, Teixeira AA. 

The West Nile virus encephalitis outbreak in the 

United States (1999-2000): from Flushing, New 

York, to beyond its borders. Ann NY Acad Sci. 

2001;951:161-171.

12. Murray KO, Mertens E, Despres P. West Nile virus 

and its emergence in the United States of America. 

Vet Res. 2010;41(6):67.

13. Hadler JL, Patel D, Bradley K, et al. National capacity for surveillance, prevention, and control of West 

Nile virus and other arbovirus infections - United 

States, 2004 and 2012. MMWR. 2014;63(13):281-

284. Available at: https://www.cdc.gov/mmwr/pre

view/mmwrhtml/mm6313a2.htm. Accessed March 

17, 2017.

14. American Mosquito Control Association. Policy 

Statement: Epidemiology and laboratory capacity grants for mosquito-borne disease surveillance 

[internet]. 2015. Available at: http://www.mosquito.

org/assets/WashingtonConf/2015/federal%20fund

ing%20issue%20paper%202015.pdf. Accessed 

February 2, 2017.

15. Womack ML, Thuma TS, Evans BR. Distribution 

of Aedes albopictus in Georgia USA. J Am Mosq

Control Assoc. 1995;11:237.

16. US Census Bureau. 2010 Census Interactive Population Search [internet]. Available at: https://www.

census.gov/2010census/popmap/ipmtext.php?fl=13. 

Accessed January 29, 2017.

17. Georgia Department of Community Health. Georgia rural county map [internet]. August 2008. 

Available at: https://dch.georgia.gov/documents/

georgia-rural-county-map. Accessed February 3, 

2017.

GEORGIA’S COLLABORATIVE APPROACH TO EXPANDING MOSQUITO SURVEILLANCE

IN RESPONSE TO ZIKA VIRUS: A CASE STUDY
January – June 2017 33

THE ARMY MEDICAL DEPARTMENT JOURNAL

18. Juliano SA. Coexistence, exclusion, or neutrality? A meta-analysis of competition between Aedes albopictus and resident mosquitoes. Isr J Ecol. 

2010;56:325-351.

19. University of Florida. Culex nigripalpus [internet]. 

Updated January 2013. Available at: http://entnem

dept.ufl.edu/creatures/aquatic/fl_sle_mosquito.

htm. Accessed February 6, 2017.

20. Goddard LB, Roth AE, Reisen WK, Scott TW. Vector competencies of California mosquitoes for West 

Nile virus. Emerg Inf Dis. 2002;8(12),1385-1391.

21. Virginia Mosquito Control Association. Coquillettidia perturbans [internet]. Available at: http://

mosquito-va.org/?page_id=437. Accessed February 6, 2017.

22. Bara JJ, Muturi EJ. Container type influences the 

relative abundance, body size, and susceptibility of 

Ochlerotatus triseriatus (Diptera: Culicidae) to La 

Crosse virus. J Med Entomol. 2015;52(3):452-460.

23. US Air Force News Service. Dobbins combats Zika 

with Georgia department of Public Health [internet]. Available at: http://www.af.mil/News/Article

Display/tabid/223/Article/925942/dobbins%ADc

ombats%ADzika%ADwith%ADgeorgia%ADdep

artment%ADof%ADpublic%ADhealth.aspx. Accessed February 6, 2017.

Authors

Dr Rustin is an Assistant Professor, Department of Epidemiology and Environmental Health Science, JiannPing Hsu College of Public Health, Georgia Southern 

University, Statesboro, Georgia.

Mr Martin is a MPH graduate student, Department of 

Epidemiology and Environmental Health Science, JiannPing Hsu College of Public Health, Georgia Southern 

University, Statesboro, Georgia.

Mr Sevilimedu is a DrPH (Biostatistics) student at the 

Jiann-Ping Hsu College of Public Health, Georgia Southern University, Statesboro, Georgia.

Mr Pandeya is a DrPH (Biostatistics) student, Department of Biostatistics, Jiann-Ping Hsu College of Public Health, Georgia Southern University, Statesboro, 

Georgia.

Dr Rochani is an Assistant Professor, Department of 

Biostatistics, Jiann-Ping Hsu College of Public Health, 

Georgia Southern University, Statesboro, Georgia.

Dr Kelly is a Public Health Entomologist with the VectorBorne & Zoonotic Diseases Team Environmental Health 

Section, Georgia Department of Public Health, Atlanta, 

Georgia.
34 http://www.cs.amedd.army.mil/amedd_journal.aspx

In 2016, Zika virus and congenital infections became 

nationally notifiable conditions in the United States.1

A total of 2,382 confirmed and probable cases of Zika 

virus (ZIKAV) with illness onset were reported to ArboNET, the US national arboviral surveillance system 

managed by the Centers for Disease Control and Prevention and state health departments, from January 1 to 

July 31, 2016.2

 In July 2016, the first locally acquired case 

of ZIKAV disease from mosquitoes was confirmed in 

the state of Florida.3 Aedes mosquitoes transmit ZIKAV, 

chikungunya virus (CHIKV), dengue virus (DENV), 

and yellow fever virus, among others. Although Aedes albopictus Skuse is thought to be a competent vector of ZIKAV,4 Ae. aegypti (L.) has been implicated as 

the primary transmitter of the virus in human populations in the ongoing outbreak in the Americas.5,6 This is 

likely the result of Ae. aegypti preferring to feed more 

frequently on humans,7,8 and being highly peridomestic 

compared to Ae. albopictus, which can inhabit more 

rural environments.9,10 The role of other mosquito species in ZIKAV transmission is either unknown, refers to 

species not present in the United States, or is controversial,11 therefore only Ae. aegypti and Ae. albopictus are 

considered here.

In this study, we concentrated on US Department of Defense (DoD) facilities, but the approach could be used for 

any area of interest. Some military facilities have longstanding mosquito surveillance programs,12 and Zika 

virus surveillance is being enhanced in the US military 

as a result of the recent threat. For example, surveillance efforts are supported by funding from the Global 

Emerging Infections Surveillance and Response section 

of the Armed Forces Health Surveillance Branch in the 

Defense Health Agency’s Public Health Division.13 According to a March 2016 DoD memo, 190 DoD installations are located in areas where mosquitoes capable of 

carrying ZIKAV occur, and increased vector monitoring will be conducted in installations in 27 states, the 

District of Columbia, Guam, and Puerto Rico.14 Four 

regional commands exist under the US Army Medical 

Command, all of which have entomological sciences divisions that conduct mosquito surveillance. Additionally, the US Air Force School of Aerospace Medicine, the 

US Navy and Marine Corps Public Health Center and 

regional Navy Environmental and Preventive Medicine 

Units, and the Navy Entomology Center of Excellence 

assist those undertaking vector surveillance or arbovirus testing.

For a military entomologist tasked with establishing and 

maintaining an Aedes spp./ZIKAV surveillance program in temperate areas that experience high mosquito 

seasonality, two important questions arise: (1) is ZIKAV 

transmission possible here?; (2) when should vector surveillance be conducted? In temperate zones like the continental United States, the answers to these questions 

may vary depending on the time of year. In this article, 

we describe an Excel-based tool that is designed to assist entomologists and other health professionals address 

these 2 questions throughout the year.

Habitat suitability models displaying potential distribution have been published for both Ae. aegypti and Ae. 

A Location-specific Spreadsheet for

 Estimating Zika Risk and Timing for

 Zika Vector Surveillance, Using US

 Military Facilities as an Example

Desmond H. Foley, PhD

David B. Pecor, BS

Abstract

Local Zika virus transmission in the United States involving one or both of the known vector species, Aedes 

aegypti and Ae. albopictus, is of major concern. To assist efforts to anticipate the risks of transmission, we 

developed an Excel spreadsheet tool that uses vector and virus temperature thresholds, remotely sensed maximum temperature, and habitat suitability models to answer the questions: “is Zika transmission likely here?” 

and “when should we conduct vector surveillance?” An example spreadsheet, updated regularly and freely 

available, uses near real-time and forecast temperature data to generate guidance, based on a novel four level 

Zika risk code, for 733 US military facilities in the 50 states, the District of Columbia, and the territories of 

Guam and Puerto Rico.
January – June 2017 35

albopictus,

15-19 as well as for ZIKAV.20-23 While these 

models often display average yearly suitability, they do 

not necessarily provide information that could be used 

for decisions about the timing of surveillance activities, 

and are global in extent rather than focused on particular areas where a surveillance program might be established. Questions about timing of mosquito monitoring 

and allocation of resources requires a consideration of 

what conditions limit adult mosquito activity and ZIKAV dissemination in the field.

Relative humidity, rainfall, drought, and wind velocity 

affect survival and behavior of mosquitoes, and therefore transmission.24 However, temperature is the most 

important ecological determinant of the development 

rate of Ae. aegypti,

25 and one of the principal determinants of Aedes survival.26 Temperature also directly affects the replication rate of arboviruses, thus affecting 

the extrinsic incubation period.27 What then do we know 

about how temperature limits Aedes and arboviruses 

such as ZIKAV?

In their review of reports published in the early 20th century, Bonne-Wepster and Brug28 presented data about 

the effects of temperature on the activity and survival of 

Ae. aegypti mosquitoes, summarized below:

 Female mosquitoes were observed to feed most 

readily between 26°C and 35°C, between 19°C and 

25°C they are slow to blood feed, and below 15°C 

to 19°C they do not feed (Marchoux et al,29 Howard 

et al,30 Connor 31).

 Female mosquitoes in Montevideo can continue to 

bite at 14°C to 15°C (Cossio32).

 Adults were observed to die when exposed to temperatures above 38°C, but Davis33 reported some 

adults surviving exposure to 40°C during 7 hours, 

and amongst another group exposed to 45°C for 2 

hours.

 At 7°C to 9°C, adult females may live up to 80 

days, and the male up to 14 days (Guiteras; Otto; 

Newman34).

 The female dies within 24 hours, if exposed to a 

temperature of 6°C (Flu35).

 Dinger et al36 reported survival of adult female 

mosquitoes kept for 6 days at 5°C.

More recently, in Saudi Arabia, Khormi et al37 found that 

the minimum temperature range of 18°C to 25°C is suitable for Ae. aegypti survival, and the survival rate increases up to 38°C. Conner38 and Wayne and Graham39 found 

that Ae. aegypti is most active at temperatures between 

15°C and 30°C, while other field and laboratory observations found survival rates from about 18°C to 38°C or 

less, based on daily or monthly minimum and maximum 

temperatures.40-42 In a study of Ae. aegypti distribution 

using the program CLIMEX, Khormi and Kumar18 set 

the limiting low temperature at 18°C, the lower optimal 

temperature at 25°C, the upper optimal temperature at 

32°C, and the limiting high temperature at 38°C. Brady 

et al16 limited their predictions of temperature suitability 

to areas with a maximum monthly temperature exceeding 13°C for Ae. albopictus and 14°C for Ae. aegypti. 

These threshold temperatures were based on previous 

studies of the observed temperatures below which biting 

and movement behaviors were impaired.42-45

Studies suggest that an increase between 14°C to 18°C 

and 35°C to 40°C can lead to higher transmission of dengue.46 Xiao et al47 found that oral infections of DENV2 

did not produce antigens in the salivary glands of Ae. albopictus kept at 18°C for up to 25 days but did produce 

antigens at 21°C. It is not known if Ae. albopictus held 

longer at the lower temperature would have disseminated 

infections, but Dohm et al48 found that Culex pipiens required 25 days at 18°C to disseminate infections of West 

Nile virus (WNV). For comparison, WNV is capable of 

replication from 14°C to 45°C.49,50 Tilston et al51 analyzed 

monthly average temperature of cities that experience 

chikungunya outbreaks and found that start and finish 

occurred when average monthly temperatures were 20°C 

or higher. At the upper temperature limit, Kostyuchenko 

et al52 found that ZIKAV is more thermally stable than 

DENV, and is also structurally stable even when incubated at 40°C, mimicking the body temperature of extremely feverish patients after virus infection. However, 

a study by Goo et al53 indicates that the thermal stability 

of some ZIKAV strains, including those involved in recent outbreaks, falls between those of DENV and WNV.

Remotely sensed temperature data is freely available 

from multiple sources as both near-real time recordings 

and forecast predictions. Combining remotely-sensed 

temperature data with predicted distributions of the vectors and virus could provide insight into when areas of 

interest are suitable for transmission and should be actively monitored. Our aim was to produce a knowledge 

product and surveillance decision tool that uses publicly 

available information about potential distribution and 

thermal requirements of the vectors and virus at US 

military facilities.

Materials and Methods

Areas of Interest

The location and boundary of US military facilities was 

obtained from the US Census Bureau’s TIGER/Line 

THE ARMY MEDICAL DEPARTMENT JOURNAL
36 http://www.cs.amedd.army.mil/amedd_journal.aspx

2015 shapefile product.54 This shapefile lists facilities in 

the continental United States, Alaska, Hawaii, Puerto 

Rico, and Guam. As some facility names comprised 

multipart polygons, these were reduced from 804 to 733 

to match the number of unique facility names, using the 

Dissolve tool in ArcMap 10.4 (ESRI, Redmond, CA). 

The centroid of each facility was selected to produce a 

shapefile of points using the Feature to Point tool (inside 

polygon option checked) of ArcMap. The georeference 

of each centroid was obtained by the Add XY Coordinates tool and joined to the points shapefile. Extraction 

of all facility centroid raster values was first obtained 

by the Extract Values to Points tool, then for polygons 

using the Zonal Statistics as Table tool, and the results 

merged. This approach was required because smaller 

polygons would not produce results using the Zonal Statistics as Table tool, which necessitated use of the raster 

data associated with the points for these facilities.

Temperature Data

To monitor temperature in near real-time, daily time averaged maps of air temperature at the surface (daytime/

ascending) were downloaded from the Giovanni 4.19 

(Released Date: 2016-04-12. Data provided by the National Air and Space Administration (NASA) Goddard 

Earth Sciences Data and Information Services Center) 

data portal at 1° spatial resolution.

55 Daily gridded temperature analyses were also collected from the National 

Oceanic and Atmospheric Administration (NOAA) US 

National Weather Service Climate Prediction Center 

(CPC).

56 Forecast temperature data was also provided 

by the CPC and the NOAA National Digital Forecast 

Database at 5 km spatial resolution.

57 For predictions 

based on monthly averages, monthly gridded climate 

data with a spatial resolution of 1 km were downloaded 

from WorldClim.58

Habitat Suitability Models

We chose the models of Ae. aegypti and Ae. albopictus 

by Kraemer et al,59 which were based on an extensively 

documented set of presence observations for each vector. We also used the habitat suitability model for ZIKAV transmission by Messina et al.21 This model used 

Zika case reports and data on temperature, precipitation, 

humidity, enhanced vegetation index, and urban versus 

rural. Both vector and virus maps are at a 5 km x 5 km 

spatial resolution. The 0.5 model suitability score was 

arbitrarily used as a presence/absence threshold.

Thresholds

The temperature suitable for activity of Ae. aegypti

and Ae. albopictus combined was estimated as 13°C

to 38°C, and 18°C to 42°C for ZIKAV. We chose to be 

conservative, using temperatures at the extremes of the 

reported suitable temperature range, and maximum 

rather than mean air temperatures.

Human Population Data

In order to more fully understand the potential impact of 

ZIKAV risk to military and nonmilitary personnel and 

their families in and around each facility, we explored 

risk in terms of human population data, with the following considerations. The flight range of Ae. aegypti

and Ae. albopictus is in the order of hundreds of meters 

only,

60,61 and each facility would differ in the average 

distance that human carriers of ZIKAV would routinely 

travel to and from each facility. Additionally, some facilities were remote, while others were adjacent to or 

enclosed within urban and suburban areas. To address 

these complications, we created a buffer of 5 km around 

all facility polygons to capture the human population 

density according to LandScan 2011.62 This was accomplished using the LandScan raster and the Buffer and 

Sum output in the Zonal Statistics as Table tools in ArcMap. A buffer of 5 km is a conservative estimate and is 

meant to give a uniform measure for each facility of the 

potential host density affected in an outbreak or vector 

control situation.

Excel-based Zika Risk Tool

A goal of this project was to display disparate data 

sources visually and in a simple and intuitive way in 

order to more effectively communicate the level of risk 

at each military facility. The risk estimation and alert 

system had to be in a format that was readily understandable and easily accessible by military users, who 

often have information system security restrictions or 

bandwidth caps. We chose MS Excel (Microsoft Corp, 

Seattle, WA), as a universal platform for performing 

calculations and reporting results. This software had 

the added advantage that the scatterplot function can be 

used to map each military facility,

63 with icons displaying various categories of risk, and using a geocorrected 

map background64 for each state. Other notable features 

that were used in the Excel risk estimation tool were 

the formula functions, conditional formatting to represent categories of numbers as different types of symbols, 

dependent dropdown lists and hyperlinks to allow users to navigate more quickly to the results of individual 

facilities, and textualized results that users can read as 

statements describing the situation and as guidance for 

vector surveillance.

Calculations Within the Excel Risk Estimation Tool

Given that the maximum temperature is available 

for a site (ie, “Temp”), the following lists an example 

A LOCATION-SPECIFIC SPREADSHEET FOR ESTIMATING ZIKA RISK AND TIMING FOR

ZIKA VECTOR SURVEILLANCE, USING US MILITARY FACILITIES AS AN EXAMPLE
January – June 2017 37

THE ARMY MEDICAL DEPARTMENT JOURNAL

sequence of tasks and their calculations, with explanations and the Excel formula (in square brackets), culminating 

in a risk rating:

1. Column A. “Was temperature suitable during period for the vector?”, ie, if the maximum was 13°C to 38°C, 

it is 1, otherwise 0 [=IF((Temp>=13)-(Temp>38),1,0)],

2. Column B. “Was temperature suitable during period for virus replication in mosquito?”, ie, if the maximum 

was 18°C to 42°C, it is 1, otherwise 0 [=IF((Temp>=18)-(Temp>42),1,0)]

3. Column C. “What is the sum of the thermal suitability values for vector (Column A) and virus (Column 

B)?”, ie, possible choices are: 0, 1, or 2.

4. Column D. “If temperature for the vectors (Column A) was within the required range, what is the model 

suitability for vector?”, ie, this was the maximum modeled suitability (0-1.00) for either Ae. aegypti or Ae. 

albopictus.

5. Column E. Score vector model suitability as 3 if ≥0.5, otherwise 2 [=IF(Column D<0.5,2,IF(Column 

D>=0.5,3))]. The 0.5 model suitability score was arbitrarily used as the cutoff for presence/absence.

6. Column F. “If temperature for the virus (Column B) was within the required range, what is the model suitability for the virus?” (0-1.00)

7. Column G. Score virus model suitability as 7 if ≥0.5, otherwise 5 [=IF(Column F<0.5,5,IF(Column 

F>=0.5,7))]. The 0.5 model suitability score was arbitrarily used as the cut-off for presence/absence.

8. Column H. “What is the ‘Combined Score’ for the interaction of temperature suitability of vector and virus, 

vector model suitability, and virus model suitability score (ie,=C×E×G)?” The use of 0, 1, and prime numbers for the component scores produces a unique semi-prime number for the product, ie, 0, 10 (=1×2×5), 14 

(=1×2×7), 15 (=1×3×5), 21 (=1×3×7), 20 (=2×2×5), 28 (=2×2×7), 30 (=2×3×5), or 42 (=2×3×7). A zero indicated that the temperature at the site was unsuitable for both vector and pathogen, so suitability scores were 

irrelevant, and combination scores were all scored zero.

9. The 9 possible Combined Scores were divided into 6 categories based on whether preconditions do not exist 

for transmission, are unsuitable for transmission, are somewhat suitable for transmission, or are suitable 

for transmission (Figure 1).

10. Column I. An Overall Zika Risk Code was established based on the Combined Score (Figure 1), and rates 

conditions as low (Blue: Code 1) to high risk (Red: Code 4). The 9 possible Combined Scores are initially 

divided according to whether temperature conditions are not suitable (Code 1) or suitable for the vectors 

and virus (Codes 2-4). Codes 2-4 are then characterized according to increasing habitat suitability, with 

Code 4 being where models predict suitable habitat for vectors and virus.

11. Action statements were constructed based on the temperature and habitat model suitability scores (Figure 

2). For example, if conditions are too cold for the development of the vectors and models predict that the 

location is unsuitable for the vectors, the Action statement would be: “Too cold or hot for vectors–surveillance unnecessary. When Temp suitable, model suggests vectors unlikely or low numbers.” Alternatively, 

if conditions are warm enough for the development of the vectors and models predict that the location is 

highly suitable for the vectors, the Action statement would be: “Temp suitable for vectors–surveillance may 

be needed. When Temp suitable, model suggests vectors likely–may need control, education, and policies 

minimizing exposure.”

Results

The Excel files comprise 12 monthly files based on average monthly maximum temperatures (suitable for 

longer term planning), and near real-time and forecast 

file, updated weekly. These files are freely available via 

the VectorMap website.

65 The tool provides risk maps 

of facilities as a continental overview (Figure 3), and 

on a state by state basis (Figure 4). Results for individual facilities are navigable via dropdown menus and 

hyperlinks (Figure 5). Statewide summary data of risk 

profile and humans potentially affected is shown in Figure 6. The temporal changes in average risk based on 

the 12 monthly files is given in Figure 7 in terms of the 

number of facilities affected (of 733) and the number of 

people within 5 km of the facilities. April to October 

was the period of greatest risk with suitable conditions 

for Zika transmission (ie, code 4) potentially affecting 

a maximum of 114 facilities in 12 states and territories, 
38 http://www.cs.amedd.army.mil/amedd_journal.aspx

and 4,546,505 people within the vicinity of these facilities, of a total of 32,811,618 within the vicinity of all 733 

facilities. The maximum number of facilities recording 

code 4 in any one month (eg, August) were: Florida (36), 

Hawaii (16), Louisiana (12), Texas (11), and Virginia (11). 

Of these, the number of people within 5 km of these 

facilities were: Texas (1,215,230), Florida (1,125,032), 

Louisiana (462,586), Virginia (409,066), and Hawaii 

(247,918).

These data may assist public health planning, and can 

be seen as an indicator of potential disease burden, or 

of people potentially benefiting from a well-informed 

vector surveillance and control program conducted on 

military facilities. Results are provided in a variety of 

symbologies and as textualized statements of how the 

factors examined may affect ZIKAV transmission, and 

recommended actions for entomologists conducting 

routine vector surveillance. The action statement textualizes the data and is designed to assist a preparedness 

posture, particularly around vector surveillance and 

control. Changes in the action statement over the year, 

for example as a result of rising temperature, can be 

used as a guide to affect changes in vector surveillance 

and control activities at particular facilities.

Comment

The Excel tool is designed to provide insights into ZIKAV transmission potential at US military facilities, 

but could be applied to other arboviruses and situations, 

such as cities,

66 tire dumps, or parks. The spreadsheet is 

flexible in that vector and virus suitability model scores, 

temperature limits, and the wording of action statements can be replaced depending on the context, and as 

new information is compiled.

For US military situations, this tool could be used in 

conjunction with the Electronic Surveillance System for 

the Early Notification of Community-based Epidemics67

or Medical Situational Awareness in Theater,68 which 

Figure 2. Action statements constructed on the basis of the temperature and habitat model suitability scores.

Figure 1. An overall Zika Risk Code based on the combined score, which rates conditions from low risk (Blue: Code 1) to high risk 

(Red: Code 4).

A LOCATION-SPECIFIC SPREADSHEET FOR ESTIMATING ZIKA RISK AND TIMING FOR

ZIKA VECTOR SURVEILLANCE, USING US MILITARY FACILITIES AS AN EXAMPLE
January – June 2017 39

THE ARMY MEDICAL DEPARTMENT JOURNAL

reports on febrile illnesses and rash in the military 

population. Coordination of result reporting through the 

Armed Forces Pest Management Board69 and VectorMap 

may also be desirable. The Navy and Marine Corps Public Health Center (NMCPHC) guide, Aedes Surveillance 

and Control Plan for U.S. Navy and Marine Corps Installations,

70 states that “each installation’s medical personnel should conduct ongoing Aedes surveillance during the mosquito season appropriate to their region and 

take preventive and responsive action to reduce disease 

risk to active duty, government employees, and family 

member populations.”70(p4) In addition, it points out that 

“OPNAVINST 6250.4C...requires all Navy and Marine 

Corps installations to have an Emergency Vector Control Plan (EVCP) for disease vector surveillance and 

control during disease outbreaks.”70(p4) The spreadsheet 

described in this study should provide an additional resource for installations as they use the NMCPHC guide 

to “...complement installation pest management plans, 

including the EVCP, as a way to assess the risk of vector 

borne diseases, and implement strategies to reduce the 

risk to personnel assigned to installations.”70(p4)

Knowing when conditions are suitable for vectors is 

crucial for monitoring the success or failure of any control program. Appendix C of the NMCPHC guide70 is 

a chart to determine the risk of infection on an installation and when to apply vector control measures. This 

4-level vector threat response plan relies on information 

about vector abundance and reports of disease transmission. We believe the Excel spreadsheet risk tool to be 

a valuable adjunct to the NMCPHC plan, as it would 

assist with defining the length of the mosquito season, 

and the judicious deployment and timing of entomological resources. Each military facility is unique and varies 

in size, function, human density, and suitable mosquito 

habitat, so not all of the 733 facilities addressed in this 

study will be at risk of mosquito-borne disease and suitable candidates for mosquito surveillance. However, in 

any case, all locations should be useful as points of reference for other nearby locations where mosquito surveillance is conducted.

Development of the Zika Risk Code in Figure 1 derived 

some inspiration from Figure 3 of Fischer et al,71 who 

combined models of vector habitat suitability with temperature categories for CHIKV replication to produce a 

matrix of climate related risk classes.

It is important to note that each data source used in this 

analysis has the potential for errors which should be considered when determining risk. For example, habitat suitability models for each vector may not be accurate for 

all areas, and may only predict average yearly suitability. 

Traits like temperature vary continuously over the surface of the earth but are effected by elevation and use of 

California military facilities

Outside temp.range for ZIKV Within temp. range for ZIKV

Outside temp. range for Zika vectors Within temp. range for Zika vectors

Figure 3. Average maximum temperature conditions for January for vectors and ZIKAV at military facilities within the contiguous 

48 states.
40 http://www.cs.amedd.army.mil/amedd_journal.aspx

multiple scale data, including weekly and monthly, and 

1 km and 1° spatial resolution data used here will result 

in important spatial and temporal variations in accuracy. 

Temperature data refers to the maximum daytime air 

temperature near the surface (averaged over various spatial resolutions) from daily data for a recent date range, 

which NASA acknowledges has limitations. Vectors can 

also seek microclimates (eg, indoors, subterranean habitats) that may be warmer or cooler than the outside temperature that is estimated by remote sensing data.

Temperatures within the suitable range may not affect 

organisms uniformly. According to Westbrook et al,72

adult female mosquitoes reared from immature stages 

at 18°C, were 6 times more likely to be infected with 

CHIKV than those reared at 32°C. Westbrook et al72

noted that climate factors, such as temperature, experienced at the larval stage (which would not be detected 

by adult trapping programs) can influence the competence of adult female mosquitoes to vector arboviruses. 

We also do not account for temperature fluctuations; according to Lambrechts et al,73 mosquitoes lived longer 

Figure 4. Thermal conditions for January and August for vectors and ZIKAV at military facilities in California. Note: unsuitable 

conditions in August in the south are due to temperatures too high for the vectors.

California military facilities

Outside temp.range for ZIKV Within temp. range for ZIKV

Outside temp. range for Zika vectors Within temp. range for Zika vectors

January August

A LOCATION-SPECIFIC SPREADSHEET FOR ESTIMATING ZIKA RISK AND TIMING FOR

ZIKA VECTOR SURVEILLANCE, USING US MILITARY FACILITIES AS AN EXAMPLE

Figure 5. An overall Zika virus risk code for near real-time and forecast periods (A) is assigned based on a combination of: 

temperature suitability for adult activity of the vectors (Ae. aegypti and Ae. albopictus) (B), modeled habitat suitability of the 

vectors (C), temperature suitability for Zika virus replication within the vectors (D), and modeled habitat suitability of Zika virus 

transmission (E). If suitable conditions exist (Zika Risk Code 4), the number of people within 5 km is shown (F) as one indication 

of the number of potential hosts in the vicinity, or the number of people potentially benefiting from facility-wide vector surveillance and control programs.

A B C D E F

Overall Zika

Risk Code

“1” (Blue)=low risk

“4” (Red)=high risk

Number of

people within

5 km with

Code 4 Zika level

Is it usually warm enough during 

Jan for Aedes aegypti/albopictus?

(Red=Yes; Green= No)

If above threshold temperature, 

what is annual % suitability for 

Aedes aegypti or albopictus?

(Red=Yes; Green=No)

Is it usually warm enough during 

January for Zika replication

in mosquito?

(Red=Yes; Green=No)

If above threshold temperature, 

what is annual % suitability

for Zika? 

(More red=more suitable)
January – June 2017 41

THE ARMY MEDICAL DEPARTMENT JOURNAL

and were more likely to become infected with DENV 

under moderate temperature fluctuations rather than under large temperature fluctuations. Thangamani et al74

and Ferreira-de-Brito et al6

 found that ZIKAV can be 

vertically transmitted in Ae. aegypti but not Ae. albopictus. This capability suggests mechanisms for the virus 

to survive in eggs that can survive for months in a dried 

dormant state during adverse conditions, for example, a 

harsh winter that would normally kill adult mosquitoes.

The approach to estimating risk levels in this project deliberately uses simplified assumptions about temperature 

and mosquito physiology and relies on published models 

to consider other drivers, which could include: precipitation; interspecific competition; and anthropogenic factors such as imported cases, built-up areas, vegetation 

indices, human behavior, and economic indices that can 

modify risk in complex and less understood ways. Other 

models could be used in place of the ones included in 

this study. For example, Samy et al23 used proxies for 

poverty and accessibility that may further increase the 

biological reality of estimates of transmission risk. As 

a risk factor for ZIKAV infections is congenital brain 

abnormalities including microcephaly,75 risk estimates 

would be greatly enhanced by quantifying the proximity 

of pregnancies to areas of interest76,77 (ie, military facilities). Among US military women, those of reproductive 

age represent the majority,78 and from 2001 to 2010, an 

average of 15,600 children were born to active component women each year.

79 Data from the Defense Medical 

Surveillance System do not include records of nonreimbursed care received at medical facilities outside of the 

military health system, adding to the difficulty in estimating numbers of potentially at-risk mothers, or those 

in the vicinity who could benefit from vector control 

programs conducted within military facilities.

The Zika virus can be imported and spread by nonvector transmission routes (eg, sexual transmission80), so it 

is recommended that a level of caution be taken when 

interpreting the data provided by this system. It is wise 

to monitor activity in surrounding facilities and any 

reputable information from other sources before acting 

on any recommendations given here. It is further recommended that the near-real time and forecast analysis should be viewed in conjunction with the monthly 

average Excel vector hazard files which uses average 

monthly maximum temperature, to gain further longerterm insights into where thermal conditions will support 

vector activity.

While this tool could be improved with higher resolution data and more nuanced models, our aim was to create an accessible and adaptable, yet useful platform for 

entomological decision-making that uses readily available data and models. New models and higher resolution 

climate data can be easily incorporated into this tool as 

they become available. Validation of the output from this 

tool using mosquito surveillance results, and obtaining 

user feedback, would be useful goals for future research.

State # Military 

facilities1

Zika Risk 

Code2-5 

(average)

# facilities where 

preconditions 

suitable for 

transmission (Zika 

Risk Code 4)5

# facilities with 

above threshold 

temperature for 

the vectors6

# facilities with 

above threshold 

temperature for 

the pathogen7

Total population 

within 5 km of 

facilities8

Total population 

within 5 km with 

suitable 

conditions for 

transmission

Alaska 20 1 0 0 0 357,044 0

Alabama 14 1 0 10 0 553,648 0

Arkansas 4 1 0 0 0 187,622 0

Arizona 11 2.19 0 10 0 556,120 0

California 100 1.71 0 79 0 6,655,724 0

Colorado 9 1 0 0 0 386,446 0

Connecticut 10 1 0 0 0 212,038 0

District of Columbia 8 1 0 0 0 842,772 0

Delaware 2 1 0 0 0 52,946 0

Florida 48 2.44 17 48 1 1,373,859 653,556

Georgia 16 1 0 9 0 1,021,113 0

Hawaii 36 3.2 16 36 1 980,370 247,918

Iowa 2 1 0 0 0 266,567 0

Summary of Vector hazard during January for U.S. Military facilities 

by State for Zika virus

Figure 6. Summary risk data for each US state (top of list shown) to assist with public health and resource allocation planning.
42 http://www.cs.amedd.army.mil/amedd_journal.aspx

Acknowledgments

We thank Jean-Paul Chrietien (Armed Forces Health 

Surveillance Branch (AFHSB)) for constructive feedback 

during the development of the Excel Risk Tool, as well as 

Jim Writer and Penny Masuoka for providing references 

and feedback useful in the development of the ZIKAV 

threshold information. We thank LTC Jeff Clark and 

MAJ Wes McCardle (former and current Walter Reed 

Biosystematics Unit Chiefs respectively) for their support during the development of this tool. We also thank 

LTC (Ret) Jason Richardson, MAJ Karl Korpal, CDR 

Fred Stell, and Dr Terry Carpenter of the Armed Forces 

Pest Management Board for their feedback. This study 

was made possible by FY2016 grant P2007_16_WR from 

the AFHSB and its Global Emerging Infections Surveillance and Response Section.

This research was performed under a memorandum of 

understanding between the Walter Reed Army Institute 

of Research and the Smithsonian Institution’s National 

Museum of Natural History, with institutional support 

provided by both organizations. The opinions contained 

herein are those of the authors and do not reflect official 

views of the supporting agencies.

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ZIKA VECTOR SURVEILLANCE, USING US MILITARY FACILITIES AS AN EXAMPLE
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A LOCATION-SPECIFIC SPREADSHEET FOR ESTIMATING ZIKA RISK AND TIMING FOR

ZIKA VECTOR SURVEILLANCE, USING US MILITARY FACILITIES AS AN EXAMPLE
January – June 2017 45

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Authors

Dr Foley is a Research Entomologist at the Walter Reed 

Biosystematics Unit, Entomology Branch, Walter Reed 

Army Institute of Research, and a Research Associate 

of the Entomology Department within the National Museum of Natural History, located at the Smithsonian Institution, Museum Support Center, Suitland, Maryland.

Mr Pecor is the VectorMap technical assistant at the 

Walter Reed Biosystematic Unit, Entomology Branch, 

Walter Reed Army Institute of Research, located at the 

Smithsonian Institution, Museum Support Center, Suitland, Maryland.

A LOCATION-SPECIFIC SPREADSHEET FOR ESTIMATING ZIKA RISK AND TIMING FOR

ZIKA VECTOR SURVEILLANCE, USING US MILITARY FACILITIES AS AN EXAMPLE

Aedes aegypti (courtesy of the CDC)

Aedes albopictus (courtesy of the CDC)
January – June 2017 47

Malaria is a major health threat to military and civilian 

personnel as well as family members deployed or residing in the Republic of Korea (ROK). Members of the 

Anopheles (Anopheles) are placed in 3 groups, namely 

the Hyrcanus Group (An. belenrae Rueda, An. kleini

Rueda, An. sinensis Wiedemann, An. pullus Yamada, 

An. lesteri Baisas and Hu, An. sineroides Yamada), the 

Barbirostris Group (An. koreicus Yamada and Watanabe), and the Lindesayi Group (An. lindesayi japonicus Yamada).1-3 Anopheles kleini and An. lesteri are the 

primary vectors of Plasmodium vivax malaria near the 

demilitarized zone (DMZ), while An. sinensis is a very 

poor vector and the status of the remaining species is 

undetermined.4,5 Laboratory studies have shown the potential for An. lesteri as an efficient vector6,7 and is considered a primary vector (as An. anthropophagus Xu and 

Feng) in parts of China.8

 In the ROK, the populations of 

An. lesteri are very low,9

 except on the northwest coastal 

islands.10 Females of An. belenrae and An. sineroides

have been found positive for P. vivax by enzyme-linked 

immunosorbent assay and polymerase chain reaction 

(PCR).9,11 Species in Barbirostris and Lindesayi Groups 

have not been implicated as malaria vectors in the ROK.9

Knowledge of the species composition of common 

Anopheles (Anopheles) in various habitats in different 

provinces is needed for defining malaria vector distributions, biosurveillance and vector control. Data are limited on the collections and habitats for the occurrence and 

relative abundance of indigenous Anopheles mosquitoes 

in the ROK.

With several human malaria cases reported annually 

from the ROK,10,12,13 there is an urgent need to accurately 

identify the potential Anopheles mosquito vectors. Misidentifications of species often lead to poor vector surveillance, misunderstanding of the epidemiology of 

disease transmission, and the development and implementation of ineffective strategies for control measures.1

Both morphological and molecular analyses are essential in the taxonomy and identifications of members of 

the Anopheles Hyrcanus Group species, particularly 

those where their geographic distribution and vector potential are better understood. During the mosquito surveillance, field identifications of live larvae and pupae 

may also be convenient and necessary, when possible.

Waxy body ornamentations have been previously reported on various insects, particularly scale insects,14

aphids,15 and heliconian caterpillar,16 and they have 

been shown to be useful for initial identifications during field surveillance efforts. Morphological variations 

in the body ornamentations for different stages of development, eg, leg paddles observed in the sabethine 

mosquitoes, Sabethes cyaneus (Fabricius)17 are important. Knowledge on mosquito body ornamentation is 

very limited, and little is known about the waxy body 

covering of larvae and pupae of Anopheles species or its 

importance in identifications.

Immature stages (both larvae and pupae) and adults of 

members of the An. (Ano.) Hyrcanus Group in the ROK 

cannot be identified with certainty using morphological 

techniques. Although there are some slight differences 

in the branching patterns of the setae in the larval head 

(setae 3, 7, 9-C), thorax (14-M) and abdomen (5, 9-II; 4, 

9-III; 4, 6-IV; 6-V),3

 they are still not very useful in performing accurate identifications of the species, thereby 

Biosurveillance and Morphological Variations

 of Larvae and Pupae of Common Malaria

 Vectors, Anopheles (Anopheles) Hyrcanus

 Group Species in the Republic of Korea

Leopoldo M. Rueda, PhD Terry A. Klein, PhD

Heung-Chul Kim, PhD Mustapha Debboun, PhD, BCE

Sung-Tae Chong, MS

Abstract

A total of 4,576 Anopheles (Anopheles) Hyrcanus Group larvae belonging to 6 species (An. belenrae, An. kleini, 

An. sinensis, An. pullus, An. lesteri, and An. sineroides) were collected from 7 different habitat types in 3 provinces of the Republic of Korea. The occurrence and relative abundance of 6 Anopheles species were noted. The 

descriptions in the article of the waxy body ornamentations or patterns of An. (Ano.) Hyrcanus Group larvae 

and pupae may be useful for rapid field species identification when conducting larval mosquito surveillance.
48 http://www.cs.amedd.army.mil/amedd_journal.aspx

requiring the need for the PCR techniques. Furthermore, 

specimens must be mounted on glass slides for microscopic morphological examinations of the branching 

patterns of setae of the head and other body parts, requiring excessive manpower and delays in identification. 

Some waxy ornamentations or patterns on the dorsal 

parts of the larval bodies were observed during the conduct of extensive field collections and surveillance of 

Anopheles mosquitoes from various parts of the ROK.

The main objectives of our study were: (1) to conduct 

a comparative survey of mosquito larvae from selected 

habitats using standardized sampling methods to obtain 

pertinent data on the occurrence and relative abundance 

of indigenous Anopheles species, and (2) to identify the 

waxy body ornamentations or patterns of Anopheles

(Anopheles) Hyrcanus Group larvae and pupae which 

may be useful for rapid field identifications during mosquito vector surveillance and control.

Materials and Methods

Immature mosquito collections that included both 

anophelines and culicines were conducted from 3 provinces (Gyeonggi, Gyeongsangbuk, and Chungcheongbuk) of the ROK (Figure 1). Selected habitats were sampled including the following: rice paddies (uncultivated 

and cultivated), irrigation ditches (concrete and nonconcrete ditches), drainage ditches (concrete and nonconcrete ditches, culvert and dammed ditches), ponds 

(including pond reservoir), stream margins (including 

stream ponds, inlets, algal mats), and other habitats (including ground pools, ground depressions, flood pools, 

rock pools, uncultivated fields, artificial containers).

As part of the mosquito biosurveillance, data were collected monthly from May 2007 to October 2007 and 

were combined over the sampling periods for each of 

the selected habitats in 3 provinces (Gyeonggi, Gyeongsangbuk, and Chungcheongbuk). The relative abundance 

of each Anopheles species in each habitat was calculated 

as the percentage of the total number of larvae recovered either per month or over the entire 6-month period.

As part of the morphological study of waxy body ornamentations, Anopheles larvae were collected from 9 locations in Gyeongsangbuk Province, including Hayangeup, Gyeongsansi (Collection nos. KSK-509, 511, 513, 

514, 520, 522, 523) and Cheongsong-eup, Jinbo-myeon 

(Collection nos. KSK-529, 533, 543, 545).

Depending on the habitats, larvae and pupae were collected using a standard larval dipper (350 ml, 13 cm diameter) or a white plastic larval tray (25 cm x 20 cm x 4 

cm) (BioQuip, Rancho Dominguez, CA). Collected live 

larvae and pupae, with intact waxy ornamentations or 

body patterns were placed in plastic Whirl-Pak bags (118 

ml, 8 cm x 18 cm) (BioQuip, Rancho Dominguez, CA) 

filled approximately 1⁄2 full with water from the collection site. The Whirl-Pak was tightly closed 

to retain air, placed in a cooler, and brought 

to the laboratory where selected specimens 

were photographed on the same day, and 

3rd and 4th instar larvae then placed and 

directly preserved in 100% ethanol (EtOH) 

for molecular identification. The remaining larvae were individually link-reared to 

adult stage as morphological voucher specimens for this work. Emergent mosquito 

adults were pinned on paper points, each 

given a unique collection number, and 

identified using diagnostic morphological 

characters. For molecular species identifications, DNA was isolated from individual 

larval mosquitoes and adults (1 or 2 legs 

per adult) by extraction procedure and direct sequencing as described in Wilkerson 

et al.12 Sequencing of An. sinensis, An. pullus, An. lesteri, An. belenrae, and An. kleini

are those of Wilkerson et al12 and Li et al18

using the primers therein. GenBank accession numbers for the above species are 

in Wilkerson et al12 and Li et al.18 Voucher 

specimens and collection records were 

BIOSURVEILLANCE AND MORPHOLOGICAL VARIATIONS OF LARVAE AND PUPAE OF COMMON MALARIA

VECTORS, ANOPHELES (ANOPHELES) HYRCANUS GROUP SPECIES IN THE REPUBLIC OF KOREA

Figure 1. Geographic locations of the 3 provinces in the Republic of Korea 

within which mosquitoe collections were conducted. Latitude (degrees)

38

37

36

39

35

34

33

Longitude (degrees)

125 126 127 128 129 130

0 45,000 90,000 180,000 Meters
January – June 2017 49

THE ARMY MEDICAL DEPARTMENT JOURNAL

deposited in the US National Museum of Natural History, Smithsonian Institution, Suitland, Maryland.

Results

Occurrence and Relative Abundance of Mosquito 

Larvae from Selected Habitats

A total of 4,576 specimens belonging to 6 species of An.

(Ano.) Hyrcanus Group species (An. belenrae, An. kleini, 

An. sinensis, An. pullus, An. lesteri, and An. sineroides) were collected monthly from May 2007 to October 

2007 from 7 general habitats by standard larval sampling method. As shown in the Table, all 6 species were 

collected from irrigation ditches, while only 5 species 

were collected from rice paddies, ponds, swamps, and 

other habitats, and only 4 species were collected from 

drainage ditches, and stream margins. In the irrigation 

ditches, An. sinensis (71.82%) was the most abundant 

species, followed by An. kleini (12.84%) and An. sineroides (9.68%). Anopheles larval population increased 

from 19.92% in June to 24.50% in August. In the rice 

paddies, An. sinensis (82.09%) had the greatest relative 

abundance followed by An. pullus (10.38%) and An. kleini (5.04%). Larval population increased from 16.03% in 

May to 23.81% in June. Except for An. lesteri, all species were collected from the rice paddies. In the pond 

reservoir, An. sinensis (54.63%) was the dominant species, followed by An. kleini (22.82%) and An.sineroides

(12.55%). Larval population increased from 15.75% in 

July to a peak of 24.28% in August. In the stream margins and swamps, respectively, An. sinensis (80.07%, 

36.86%) ranked first in abundance followed by An. kleini (13.07%, 27.78%) and An. pullus (5.85%, 30.38%). 

Larval population increased from 23.04% and 12.08% 

in July to 26.37% and 29.31% in August. In the drainage 

ditches, An. sinensis (51.98%) was the dominant species, 

followed by An. pullus (20.63%) and An. kleini (9.33%). 

Larval population increased from 15.38% in June to a 

peak of 35.90% in August. In all sampled habitats, An.

sinensis had the highest relative abundance in all habitats, and its larvae were found in all 7 habitats similar 

to An. kleini, An. pullus, and An. sineroides. Anopheles

lesteri larvae were only collected from the irrigation 

ditches and swamps, while An. belenrae larvae were 

only found in rice paddies, irrigation ditches, and ponds. 

Rueda et al2

 reported the associations of An. (Ano.) with 

other species of Aedes and Culex from different habitats 

in the ROK.

Morphological Descriptions of 5 Common Anopheles

(Anopheles) Species, with Emphasis on Waxy 

Ornamentations or Body Patterns, Based on 

Korean Specimens

1. An. belenrae, Larva: Body coloration is usually 

green or light reddish brown. Thorax has dorsal parts 

almost covered with white waxy materials, primarily 

along the anterior and lateral edges. It usually has 

a slightly pointed waxy ornamentation extending up 

to the anterior tip of the dorsum of abdominal segment I (Figure 2; collection nos. KSK-545-1001L, 

KSK-545-1008L, KSK-545-1011L). Abdomen has 2 

distinct white waxy ornamentations on the dorsal 

parts of segments III and V. Abdominal segment III 

has a dorsal part with white waxy ornamentations 

of various shapes, including barrel shape (Figure 

2A), diamond shape (Figure 2B), and chalice-shape 

(Figure 2C). Abdominal segment V has a dorsal part 

with a white waxy ornamentation that is almost similar to that one on abdominal segment I (Figure 2).

Pupa: No live specimen was available to examine.

2. An. kleini, Larva: Body coloration is usually reddish brown to dark brown. Thorax has dorsal parts 

almost covered with white waxy materials, primarily along the anterior half and narrowly extending 

towards the anterior edge of abdominal segment I. 

The tip of this extension is usually blunt (Figure 3B, 

3C) or rarely pointed (Figure 3A). Abdomen has one 

distinct white waxy ornamentation, barrel-shaped, 

on the dorsal part of segment III (Figure 3; collection nos. KSK-511-1004L, KSK-523-1004L, KSK545-1005L), and nothing on abdominal segment V.

Pupa: Abdominal segments III and IV have dorsal parts with white waxy ornamentations of various shapes, including Aladdin lamp shape (Figure 

4A, collection no. KSK-511-1006P), blown tubular 

glass shape (Figure 4B, collection no. KSK-513-

1004P) and cork crown shape (Figure 4C, collection no. KSK-514-1008P; Figure 5A, collection no. 

KSK-522-1004P). Figure 2. Mosquito larvae of Anopheles belenrae, dorsal view, showing 

waxy ornamentations of the thorax and abdomen (arrows).

BE(KSK-545-1001L)

A

BE(KSK-545-1008L)

B

BE(KSK-545-1011L)

C
50 http://www.cs.amedd.army.mil/amedd_journal.aspx

BIOSURVEILLANCE AND MORPHOLOGICAL VARIATIONS OF LARVAE AND PUPAE OF COMMON MALARIA

VECTORS, ANOPHELES (ANOPHELES) HYRCANUS GROUP SPECIES IN THE REPUBLIC OF KOREA

Relative abundance of Anopheles (Anopheles) larvae on various habitats in 3 provinces (Geonggi, Gyeongsangbuk, 

Chungcheongbuk) of the Republic of Korea (May 2007 - October 2007) [part 1 of 2].

Relative Abundance (%)a

Habitat and Species May June July August Sepember October 6-month 

Mean

Total

Collected (n)

Rice paddies (n=111)b

An. belenrae 1.34 0.00 0.00 0.57 0.00 0.62 0.42 0.38 (7)

An. kleini 3.01 12.61 10.00 1.98 2.62 0.00 5.04 6.06 (113)

An. lesteri 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 (0)

An. pullus 54.18 4.50 0.88 1.98 0.75 0.00 10.38 10.4 (194)

An. sinensis 33.44 82.88 88.82 95.47 96.25 95.68 82.09 81.45 (1519)

An. sineroides 8.03 0.00 0.29 0.00 0.37 3.70 2.07 1.72 (32)

Total collected (n) 299 444 340 353 267 162 (1865)

Irrigation ditches (n=84)c

An. belenrae 11.11 0.00 0.55 0.78 1.27 0.00 2.29 1.24 (13)

An. kleini 29.17 2.87 18.78 13.62 11.46 1.16 12.84 11.06 (116)

An. lesteri 0.00 0.96 0.00 0.00 0.00 0.58 0.26 0.29 (3)

An. pullus 4.17 6.70 2.21 1.95 2.55 1.16 3.12 3.05 (32)

An. sinensis 51.39 61.72 78.45 81.71 60.51 97.11 71.82 74.45 (781)

An. sineroides 4.17 27.75 0.00 1.95 24.20 0.00 9.68 9.91 (104)

Total collected (n) 72 209 181 257 157 173 (1049)

Drainage ditches (n=18)d

An. belenrae 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 (0)

An. kleini 0.00 0.00 14.29 25.00 16.67 0.00 9.33 12.82 (5)

An. lesteri 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 (0)

An. pullus 0.00 16.67 7.14 0.00 0.00 100.00 20.63 12.82 (5)

An. sinensis 0.00 83.33 78.57 75.00 75.00 0.00 51.98 71.79 (28)

An. sineroides 0.00 0.00 0.00 0.00 8.33 0.00 1.39 2.56 (1)

Total collected (n) 0 6 14 4 12 3 (39)

Ponds (n=61)e

An. belenrae 0.16 0.00 0.00 0.99 1.10 0.00 3.86 1.44 (12)

An. kleini 50.00 16.16 41.22 16.83 12.15 0.55 22.82 17.55 (146)

An. lesteri 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 (0)

An. pullus 13.16 8.08 1.53 6.93 3.31 3.87 6.15 5.05 (42)

An. sinensis 13.16 8.08 57.25 75.25 78.45 95.58 54.63 66.71 (555)

An. sineroides 2.63 67.68 0.00 0.00 4.97 0.00 12.55 9.25 (77)

Total collected (n) 38 99 131 202 181 181 (832)

Stream margins (n=32)f

An. belenrae 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 (0)

An. kleini 66.67 4.76 2.06 0.90 4.05 0.00 13.07 2.14 (9)

An. lesteri 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 (0)

An. pullus 33.33 0.00 0.00 0.00 0.00 1.74 5.85 0.71 (3)

An. sinensis 0.00 95.24 97.94 99.10 95.95 92.17 80.07 95.49 (402)

An. sineroides 0.00 0.00 0.00 0.00 0.00 6.09 1.01 1.66 (7)

Total collected (n) 3 21 97 111 74 115 (421)

Notes:

a. Relative abundance means for 6 months are based on the total number of Anopheles larvae collected during the period.

b. Rice paddies, vacant or with rice plants

c. Irrigation ditches include concrete and nonconcrete ditches

d. Drainage ditches include concrete and nonconcrete ditches, culverts, dammed ditches

e. Ponds include pond reservoirs

f. Stream margins include stream ponds, stream inlets, algal mats
January – June 2017 51

THE ARMY MEDICAL DEPARTMENT JOURNAL

3. An. pullus, Larva: Body coloration is usually green 

or brown. Thorax has dorsal parts almost covered 

with white waxy materials, primarily along the middle part, and narrowly extending towards the anterior 

edge of abdominal segment I. The tips of this waxy 

extension are usually blunt (Figure 6A, 6C; collection 

nos. KSK-543-1005L, KSK-543-1013L) or seldom 

pointed (Figure 6B, collection no. KSK-543-1006L). 

Abdomen has one distinct white waxy ornamentation, barrel-shaped, on dorsal part of segment III 

(Figure 6), and nothing on the abdominal segment V.

Pupa: Abdominal segments III and IV have dorsal parts with white waxy ornamentations of various shapes, including Aladdin lamp shape (Figure 

7A, 7B; collection nos. KSK-514-1001P, KSK-523-

1003P), and pear shape (Figure 7C, collection no. 

KSK-520-1001P).

4. An. sinensis, Larva: Body coloration is usually 

green, reddish brown, or dark brown. Thorax has 

dorsal parts almost completely covered with white 

or creamish colored waxy materials, primarily along 

the anterior half, and narrowly extending towards the 

anterior edge of abdominal segment I. The tip of this 

extension is usually blunt (Figure 8A, 8B; collection 

nos. KSK-543-1003L, KSK-545-1007L). Abdomen 

has one distinct white or creamish waxy ornamentation, barrel-shaped or pear-shaped, on dorsal part 

of segment III (Figure 8A, 8B), and usually lacking 

ornamentation on abdominal segment V. It is very 

rare that dorsal parts of the abdomen have 2 white 

waxy ornamentations. Greenish larvae usually have 

no or very light waxy ornamentations on the body.

Pupa: Cephalothorax has a cup-shaped white waxy 

ornamentation on dorsal part. Abdominal segments 

I-V have median longitudinal white waxy ornamentations on dorsal parts (Figure 5B, 5C; collection nos. 

KSK-522-1001P, KSK-523-1002P).

5. An. lesteri, Larva: Body coloration is usually green 

or dark brown. Thorax has dorsal parts almost covered with white waxy materials, and approximately 

Relative abundance of Anopheles (Anopheles) larvae on various habitats in 3 provinces (Geonggi, Gyeongsangbuk, 

Chungcheongbuk) of the Republic of Korea (May 2007 - October 2007) [part 2 of 2].

Relative Abundance (%)a

Habitat and species May June July August Sepember October 6-month 

Mean

Total

Collected (n)

Swamps (n=13)

An. belenrae 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 (0)

An. kleini 83.33 4.65 35.00 41.24 2.44 0.00 27.78 20.24 (67)

An. lesteri 0.00 0.00 5.00 1.03 7.32 0.00 2.22 1.81 (6)

An. pullus 16.67 90.70 10.00 2.06 2.44 1.02 20.48 14.80 (49)

An. sinensis 0.00 0.00 50.00 48.45 39.02 83.67 36.86 49.85 (165)

An. sineroides 0.00 4.65 0.00 7.22 48.78 15.31 12.66 13.29 (44)

Total collected (n) 12 43 40 97 41 98 (331)

Other habitats (n=22)b

An. belenrae 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 (0)

An. kleini 0.00 0.00 0.00 6.25 0.00 0.00 1.04 2.56 (1)

An. lesteri 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 (0)

An. pullus 0.00 0.00 8.33 0.00 0.00 0.00 1.39 2.56 (1)

An. sinensis 0.00 0.00 83.33 81.25 100.00 50.00 52.43 76.92 (30)

An. sineroides 100.00 100.00 8.33 12.50 0.00 50.00 45.14 17.95 (7)

Total collected (n) 2 1 12 16 6 2 (39)

Notes:

a. Relative abundance means for 6 months are based on the total number of Anopheles larvae collected during the period.

b. Other habitats including ground pools, ground depressions, flood pools, rock pools, uncultivated fields, artificial containers

Figure 3. Mosquito larvae of Anopheles kleini, dorsal view, showing 

waxy ornamentations of the thorax and abdomen (arrows).

KL(KSK-511-1004L)

A

KL(KSK-523-1004L)

B

KL(KSK-545-1005L)

C
52 http://www.cs.amedd.army.mil/amedd_journal.aspx

cone-shaped along the median parts. It usually 

has a slightly pointed waxy ornamentation extending up to the anterior tip of the dorsum of 

abdominal segment I (Figure 8C, collection no. 

KSK-545-1009L). Abdomen has 2 distinct white 

waxy ornamentations on dorsal parts of segments 

III and V. Both ornamentations are barrel-shaped.

Pupa. No live specimen was available to examine.

Comment

The seasonal occurrence and relative abundance of 

Anopheles larvae varied from one type of habitat to 

another. These variations may be due to different 

environmental conditions found in most habitats. 

Although Anopheles (Anopheles) larvae and pupae 

have been previously described based on slidemounted larval and pupal specimens,2,3,19 little is 

known about the waxy ornamentation or body covering patterns of these immature mosquitoes in nature. In spite of detailed morphological descriptions 

of the larvae and pupae in various publications,2,3,19

identifications keys are often difficult to construct 

due to much variation of supposed diagnostic characters, including position, types, forms, and number 

of branches of setae and other body parts. During 

intensive field surveillance, the use of waxy body 

ornamentations of the larvae may be useful characters for initial identifications of some Anopheles

species, but there is still a need for larval rearing 

to obtain emerged adults. Available adult identification keys as well as PCR and DNA sequencing for 

larvae, pupae, and adults may be used to ascertain 

species identifications. The shapes or patterns of 

waxy body ornamentations are also not very distinct in first-instar and second-instar larvae, unlike 

third-instar and fourth-instar larvae of Anopheles

mosquitoes. More specimens from the field, as well 

as from laboratory reared materials are required to 

further study the morphological variations or body 

patterns based on body ornamentations as diagnostic characters, in the mosquito larvae and pupae.

Acknowledgments

We thank Ms Soo-Yun Kim and Ms J. Stoffer for assistance in finalizing the photographs of mosquito larvae and pupae, and B. P. Rueda and C. R. Summers 

for providing valuable comments and suggestions to 

improve the manuscript.

This research was performed under a memorandum of 

agreement between the Walter Reed Army Institute of 

Research and the Smithsonian Institution, with institutional support provided by both organizations.

The opinions contained herein are those of the authors 

and do not reflect official views of the supporting 

agencies.

BIOSURVEILLANCE AND MORPHOLOGICAL VARIATIONS OF LARVAE AND PUPAE OF COMMON MALARIA

VECTORS, ANOPHELES (ANOPHELES) HYRCANUS GROUP SPECIES IN THE REPUBLIC OF KOREA

KL(KSK-511-1006P)

A

KL(KSK-514-1008P)

C

KL(KSK-513-1004P)

B

Figure 4. Mosquito pupae of Anopheles kleini, dorsal view, showing waxy 

ornamentations of the abdomen (arrow).

KL(KSK-522-1004P)

A

SI(KSK-522-1001P)

B

SI(KSK-523-1002P)

C

Figure 5. Mosquito pupae of Anopheles kleini (A), and An. sinensis (B,C), 

dorsal view, showing waxy ornamentations of the abdomen (arrow).

PU(KSK-543-1006L)

B

PU(KSK-543-1013L)

C

PU(KSK-543-1005L)

A

Figure 6. Mosquito larvae of Anopheles pullus, dorsal view, showing 

waxy ornamentations of the thorax and abdomen (arrows).
January – June 2017 53

THE ARMY MEDICAL DEPARTMENT JOURNAL

References

1. Rueda LM. Two new species of Anopheles

(Anopheles) Hyrcanus group (Diptera: Culicidae) from the Republic of South Korea. Zootaxa. 2005;941:1-26. http://www.mapress.com/

zootaxa/2005f/zt00941.pdf.

2. Rueda LM, Kim HC, Klein T, et al. Distribution 

and larval habitat characteristics of Anopheles

Hyrcanus Group and related mosquito species 

(Diptera: Culicidae) in South Korea. J Vector Ecol.

2006;31:199-206.

3. Tanaka K, Mizusawa K, Saugstad ES. A Revision of the Adult and Larval Mosquitoes of Japan 

(including the Ryukyu Archipelago and the Ogasawara Islands) and Korea (Diptera: Culicidae). 

Gainesville, FL: American Entomological Institute; 1979.

4. Lee WJ, Klein TA, Kim HC, et al. Anopheles kleini, An. pullus, and An. sinensis: potential vectors 

of Plasmodium vivax in the Republic of Korea. J

Med Entomol. 2007;44:1086-1090.

5. Klein TA, Kim HC, Lee WC, et al. Reemergence, 

persistence, and surveillance of vivax malaria and 

its vectors in the Republic of Korea. In: Robinson WK, Bajoni D, eds. Proceedings of the Sixth

International Conference on Urban Pests Hungary, Budapest, Hungary. 2008;325-351. Available at: http://www.icup.org.uk/icupsearchres.

asp?snum=0. Accessed February 8, 2017.

6. Joshi D, Choochote W, Park MH, et al. The susceptibility of Anopheles lesteri to infection with 

Korean strain of Plasmodium vivax. Malaria J.

2009;8:42.

7. Joshi D, Kim JY, Choochote W, Park MH, Min GS. 

Preliminary vivax malaria vector competence for 

three members of the Anopheles hyrcanus group 

in the Republic of Korea. J Am Mosq Control Assoc. 2009;27:312-314.

8. Rueda LM, Zhao T, Yajun Ma, et al. Updated distribution records of the Anopheles (Anopheles) 

hyrcanus species-group (Diptera: Culicidae) in 

China. Zootaxa. 2007;1407:43-55.

9. Rueda LM, Brown TL, Kim HC, et al. Species 

composition, larval habitats, seasonal occurrence 

and distribution of potential malaria vectors and 

associated species of Anopheles (Diptera: Culicidae) from the Republic of Korea. Malaria J.

2010;9:55-65.

10. Foley DH, Klein TA, Lee IY, et al. Mosquito species composition and Plasmodium vivax infection 

rates on Baengnyeong-do (Island), Republic of Korea. Korean J Parasitol. 2011;49:313-316.

11. Rueda LM, Li C, Kim HC, Klein TA, Foley DH, 

Wilkerson RC. Anopheles belenrae (Diptera: Culicidae), a potential vector of Plasmodium vivax in 

the Republic of Korea. J Am Mosq Control Assoc.

2010;26:430-432.

12. Wilkerson RC, Li C, Rueda LM, Kim HC, Klein TA, 

Song GH, Strickman D. Molecular confirmation of 

Anopheles (Anopheles) lesteri from the Republic of 

South Korea and its genetic identity with An. (Ano.) 

anthropophagus from China (Diptera: Culicidae). 

Zootaxa. 2003;378:1-14. Available at: http://www.

mapress.com/zootaxa/2003f/zt00378.pdf. 

13. Foley DH, Klein TA, Kim HC, Brown T, Wilkerson RC, Rueda LM. Validation of ecological 

niche models for potential malaria vectors, Anopheles (Anopheles) (Diptera: Culicidae), in the Republic of Korea. J Amer Mosq Control Assoc. 

2010;26:210-213.

Figure 7. Mosquito pupae of Anopheles pullus, dorsal view, showing 

waxy ornamentations of the abdomen (arrows).

PU(KSK-514-1001P)

A

PU(KSK-520-1001P)

C

PU(KSK-523-1003P)

B

Figure 8. Mosquito larvae of Anopheles sinensis (A,B), and An. lesteri

(C), dorsal view, showing waxy ornamentations of the thorax and abdomen (arrows).

SI(KSK-545-1007L)

B

LE(KSK-545-1009L)

C

SI(KSK-543-1003L)

A
54 http://www.cs.amedd.army.mil/amedd_journal.aspx

14. Futch SH, McCoy CW, Childers CC. A guide to 

scale insect identification. Gainsville, FL: University of Florida Institute of Food and Agricultural 

Sciences; 2015:1-5. Extension Publication #HS-817. 

Available at: http://edis.ifs.ufl.edu/ch195. Accessed 

February 2, 2017.

15. Labanoski G, Soika G. Aphids (Hemiptera, Aphidoidea) on trees and shrubs belonging to Betulacea 

family in ornamental plant nurseries. Aphid and

Other Hemipterous Insects. 2011;17: 81-97.

16. Barao KR, Silva DS, Moreira GRP. External morphology of the immature stages of Neotropical heliconians: X. Heliconius sara apseudes (Lepidoptera, Nymphalidae, Heliconiinae). Iheringia Ser

Zool. 2015;105:523-533. Available at: http://dx.doi.

org/10.1590/1678-476620151054523533.

17. South SH, Arnqvist G. Morphological variation 

of an ornament expressed in both sexes of the 

mosquito Sabethes cyaneus. Evolution Ecol Res.

2009;11:1-21.

18. Li C, Lee JS, Groebner JL, Kim HC, Klein TA, 

O’Guinn ML, Wilkerson RC. A newly recognized 

species in the Anopheles Hyrcanus Group and molecular identification of related species from the 

Republic of South Korea (Diptera: Culicidae). Zootaxa. 2005;939:1-8.

19. Harrison BA, Scanlon JE. Medical entomology 

studies - II. The subgenus Anopheles in Thailand (Diptera: Culicidae). Contr Am Entomol Inst.

1975;12:1-307.

Authors

Dr Rueda is an Adjunct Scientist of the Smithsonian Institution and former Research Entomologist and Chief 

of the Walter Reed Biosystematics Unit, Entomology 

Branch, Walter Reed Army Institute of Research, located at the Smithsonian Institution, Museum Support 

Center, Suitland, Maryland.

Dr Kim is a Research Entomologist, 5th Medical Detachment/Medical Department Activity-Korea, 65th Medical 

Brigade, Republic of Korea.

Mr Chong is an Entomologist, 5th Medical Detachment/

Medical Department Activity-Korea, 65th Medical Brigade, Republic of Korea.

Dr Klein is the senior Entomologist, Medical Department Activity-Korea, 65th Medical Brigade, Unit 15281, 

Republic of Korea.

Dr Debboun is the Director of the Mosquito & Vector 

Control Division, Harris County Public Health, Houston, 

Texas.

BIOSURVEILLANCE AND MORPHOLOGICAL VARIATIONS OF LARVAE AND PUPAE OF COMMON MALARIA

VECTORS, ANOPHELES (ANOPHELES) HYRCANUS GROUP SPECIES IN THE REPUBLIC OF KOREA
January – June 2017 55

Human Chagas disease is caused by the protozoan parasite Trypanosoma cruzi, acquired primarily through 

contact with infected excreta of triatomine insects 

(known as “kissing bugs”). Although vector-borne 

transmission predominates, humans can also become 

infected congenitally, orally through contaminated food 

or beverages, or hematogenously through blood transfusion or organ transplantation.1

 During the first 4-8 weeks 

of infection, considered the acute phase, symptoms are 

usually mild, nonspecific, or unappreciable; potentially 

fatal myocarditis or meningoencephalitis occur rarely. 

Approximately 70% to 80% of infected persons enter 

a chronic indeterminate phase, characterized by lifelong infection without symptoms, electrocardiographic 

changes, and radiographic evidence of disease. The 

remaining 20% to 30% develop chronic disease, often 

presenting years or decades after infection as clinical 

cardiac and/or digestive disease.2

With approximately 5 to 8 million people infected worldwide,1,3 Chagas is classified by the World Health Organization as one of the most important neglected tropical 

diseases.1

 There are an estimated 240,000 prevalent cases in the United States among immigrants from endemic 

areas of Latin America.4

 Fewer than 30 locally-acquired 

infections have been reported in the United States since 

1955,5

 when a resident of Corpus Christi, Texas, became 

the first documented autochthonous case in the country.6

In 2013, the Texas Department of State Health Services 

added Chagas disease to the state’s Notifiable Conditions 

list, which requires the reporting of confirmed and suspected human cases to local or regional health departments. Twelve autochthonous human infections were 

confirmed in Texas during the first 2 years of mandatory 

reporting,7

 one of which was associated with left ventricular dysfunction.8

Case Report

In October 2016, an 18-year-old US Air Force trainee 

screened positive for T cruzi infection when he donated 

blood at Joint Base San Antonio (JBSA), Texas. Blood 

from all first-time donors at the JBSA-Lackland donation center is screened for T cruzi with an enzymelinked immunosorbent assay from Ortho-Clinical Diagnostics. Per standard protocol, he was referred to the 

installation’s trainee health clinic, where he was found 

to have normal vital signs and an unremarkable physical 

exam. He reported being in excellent health and had not 

experienced any recent chest pain, shortness of breath, 

dizziness, or gastrointestinal symptoms.

A chemiluminescence immunoassay and enzyme strip 

assay (Abbott Laboratories) were ordered and found 

to be positive for anti-T cruzi IgG antibodies. A whole 

blood sample was sent to the Reference Diagnostic Laboratory at the Centers for Disease Control and Prevention for further testing. An enzyme immunoassay was 

reactive and an immunoblot of trypomastigote excretedsecreted antigens was positive, confirming the diagnosis. The patient was notified of these results and referred 

to the Department of Infectious Disease at San Antonio 

Military Medical Center (SAMMC) for further evaluation and treatment.

After notifying the patient of his laboratory results, Infectious Disease conducted a 12-lead electrocardiogram 

A Case of Chagas Cardiomyopathy

 Following Infection in South Central Texas

Maj Bryant J. Webber, USAF, MC Maj Kelvin N. Bush, USAF, MC

Lt Col Edward J. Wozniak, TXSG, MRC Maj Matthew C. Wilson, USAF, MC

CPT David Chang, MC, USA LTC James A. Watts, MC, USA

Lt Col Heather C. Yun, USAF, MC

Abstract

Between 5 and 8 million people globally are infected with Trypanosoma cruzi, the causative parasitic agent of 

Chagas disease. The vast majority of incident infections originate in pockets of Latin America where domestic 

vector-borne transmission cycles are more common. Since 1955, when the first locally-acquired case was reported, fewer than 30 autochthonous cases have been documented in the United States. We describe the case of 

an 18-year-old US Air Force trainee, a native Texan with no travel history beyond the continental United States, 

who screened positive for T cruzi infection on blood donation and was subsequently found to have chronic Chagasic cardiomyopathy. This is the first documented case of Chagas disease in a US military trainee and one of 

the first known autochthonous cases of Chagasic cardiomyopathy in a Texas resident. Diagnostic, therapeutic, 

and military implications are discussed.
56 http://www.cs.amedd.army.mil/amedd_journal.aspx

(ECG), which demonstrated normal sinus rhythm, firstdegree atrioventricular block, and left anterior hemiblock with right bundle branch block. This prompted 

referral to the Division of Cardiology at SAMMC. Cardiovascular physical exam was benign with normal heart 

sounds, normal jugular venous pressure, normal apical 

impulse, and no evidence of congestive heart failure. A 

battery of noninvasive tests was performed in order to 

assess for common cardiac manifestations of Chagas disease, including, but not limited to, left ventricular dilatation and dysfunction, wall motion abnormalities, aneurysms, diastolic dysfunction, pathologic bradyarrhythmias and tachyarrhythmias, and ischemic heart disease.9

The majority of tests were within normal limits. Chest 

x-ray showed no evidence of cardiomegaly. Holter monitoring was negative for any pathologic dysrhythmias. 

Transthoracic echocardiogram demonstrated normal diastolic, valvular, and global systolic function. Exercise 

testing with Bruce protocol established no exercise-induced arrhythmias, ischemic electrical changes, or anginal symptoms. Cardiopulmonary exercise testing found 

an appropriate VO2max, early anaerobic threshold, and 

normal VE/VCO2 slope, consistent with a subclinical 

reduction in exercise capacity with preserved ventilatory efficiency. Cardiac magnetic resonance imaging 

confirmed the diagnosis of early heart disease, demonstrating left ventricular cavity dilation with preserved 

global systolic function (ejection fraction of 76%); the 

imaging was otherwise normal with no wall motion abnormalities, late gadolinium enhancement, abnormal T1 

relaxation, or myocardial edema on T2 weighted images.

Given his exposure history, serologic findings, abnormal 

ECG, and left ventricular cavity dilation, the patient was 

determined to have chronic Chagasic cardiomyopathy. 

Per the Brazilian Consensus Classification and American College of Cardiology/American Heart Association 

classification schemes, he was classified as Stage B1 

and Stage B, respectively,10,11 and at low risk for cardiac 

death according to 2 validated risk calculators.11,12 Since 

cardiomyopathy is a disqualifiable condition for accession into the US military,13 the patient was processed for 

medical discharge from training. Infectious Disease advised the patient to complete a 60-day regimen of oral 

benznidazole,14 but he declined. He was strongly encouraged to seek follow-up in the civilian health care 

sector and to notify household contacts that they should 

be screened for Chagas disease.15

Public health personnel interviewed the patient to facilitate case reporting to the Texas Department of State 

Health Services. The patient was raised on a ranch in 

south central Texas and had never traveled outside the 

continental United States. He camped occasionally near 

his home but never hunted or skinned animals. When 

shown a display case with triatomine insects of various 

species and at different stages of development, the patient immediately recognized them, saying they “were 

all over the place” on the ranch, including within the 

residence. He did not recall ever receiving a bite. A 

number of reservoir animals were also present on the 

property, including cats, dogs, raccoons, and armadillos. The patient was not aware of any relatives having 

Chagas disease, although he was adopted at a young age 

and had no knowledge of his biological mother. He had 

never received a blood transfusion. A week before his 

blood donation, he had spent 5 days and 4 nights on the 

JBSA Medina Training Annex for a field training exercise, during which he slept in a permethrin-treated bed 

net and reported no known insect bites.

Comment

Although neither congenital acquisition nor vectorborne acquisition during military training can be definitively ruled out, this patient was likely infected with T 

cruzi while growing up on a ranch in south central Texas. Ecologic modeling has predicted that this region of 

the United States is at increased risk for autochthonous 

Chagas disease.16 Situated at the interface of tropical and 

temperate biomes, south central Texas has a number of 

environmental and cultural factors that may facilitate 

human exposure to T cruzi: a diverse array of wildlife 

reservoirs and indigenous triatomine species; the popularity of high-risk outdoor activities, especially hunting 

and camping; and the presence of scattered colonias 

(impoverished, primarily Hispanic communities). As 

compared to modern urban and suburban houses, poorly 

constructed ranches, cabins, and colonias are more susceptible to colonization by triatomine insects and wildlife reservoirs, thus increasing the likelihood of human 

exposure to infected vectors.17

The southern United States is inhabited by 11 recognized species of triatomine insects, listed in the Table, 

most of which are competent T cruzi vectors and likely 

to be involved in enzootic transmission cycles among 

indigenous wildlife reservoirs.18 All species may exist as 

nest parasites and feed on a variety of vertebrate hosts. 

Competence for transmitting the parasite to humans is 

affected by environmental distribution, dispersal capacity, feeding and defecation behaviors, and ability to invade human domiciles, attributes of which vary according to species.18-20

In south central Texas, Triatoma gerstaeckeri insects have been found to enter human dwellings and 

to feed upon humans and domestic animals.21,22 This 

A CASE OF CHAGAS CARDIOMYOPATHY FOLLOWING INFECTION IN SOUTH CENTRAL TEXAS
January – June 2017 57

THE ARMY MEDICAL DEPARTMENT JOURNAL

medium-to-large triatomine species inhabits much of 

the Edwards Plateau and South Texas Brush Country 

between the 96th and 103rd parallels, the southeastern 

corner of New Mexico, and northeastern Mexico.23 The

T cruzi infection rate of this species may exceed 60% 

in south central Texas,21,22 and adult insects often have 

detectable human blood in their midgut.24

Although the case patient was likely infected prior to arrival at JBSA, this report highlights the risk for autochthonous Chagas disease in the southern United States 

and underscores the importance of preventing Chagas 

and other vector-borne diseases while training in endemic areas. In order to decrease vector habitats, engineering controls should focus on reducing vegetation around 

military field sites to the maximum extent possible 

without disrupting the training mission. Administrative 

controls emphasizing site cleanliness should help minimize the population of woodrats, an important reservoir 

animal.25 Finally, the 4 components of optimal personal protective measures should be meticulously used: a 

properly-worn field uniform (sleeves rolled down, wrist 

openings secured, undershirt tucked into the pants, and 

pant legs tucked into the boots); permethrin treatment of 

the uniform blouse and pants; the application of either 

DEET*-based (20% to 40% concentration) or picaridin†-

based (20% concentration) insect repellent to exposed 

skin; and sleeping in a permethrin-treated bed net.26 Finally, diligent public health surveillance and health care 

provider education for Chagas disease are warranted.

References

1. Rassi A Jr, Rassi A, Marin-Neto JA. Chagas disease. Lancet. 2010;375(9723);1388-1402.

2. Bern C. Chagas’ disease. N Engl J Med. 

2015;373(5):456-466.

3. World Health Organization. Chagas disease in Latin 

America: an epidemiological update based on 2010 

estimates. Wkly Epidemiol Rec. 2015;90(6):33-43.

4. Manne-Goehler J, Umeh CA, Montgomery SP, 

Wirtz VJ. Estimating the burden of Chagas disease in the United States. PLoS Negl Trop Dis. 

2016;10(11):e0005033.

5. Montgomery SP, Parise ME, Dotson EM, Bialek SR. What Do We Know About Chagas Disease in the United States?. Am J Trop Med Hyg. 

2016;95(6):1225-1227.

6. Woody NC, Woody HB. American Trypanosomiasis (Chagas’ disease): first indigenous case in the 

United States. JAMA. 1955;159(7):676-677.

7. Texas Department of State Health Services. Chagas disease [internet]. Available at: https://www.

dshs.texas.gov/WorkArea/linkit.aspx?LinkIdentifi

er=id&ItemID=22929. Accessed January 10, 2017.

8. Garcia MN, Murray KO, Hotez PJ, et al. Development of Chagas cardiac manifestations among Texas blood donors. Am J Cardiol. 2015;115(1):113-117.

9. Nunes MA, Dones W, Morillo CA, Encina JE, Ribeiro AL. Chagas disease: an overview of clinical 

and epidemiological aspects. J Am Coll Cardiol. 

2013;62:767-776.

10. Acquatella H. Echocardiography in Chagas heart 

disease. Circulation. 2007;115:1124-1131.

11. de Souza AC, Salles G, Hasslocher-Moreno AM, et 

al. Development of a risk score to predict sudden 

death in patients with Chaga’s heart disease. Int J 

Cardiol. 2015;187:700-704.

*N,N-diethyl-meta-toluamide

†Hydroxyethyl isobutyl piperidine carboxylate

Triatoma in the United States.

Genus and species (subspecies) Discoverer Photograph

Paratriatoma hirsuta Barber

Figure 1 Triatoma gerstaeckeri Stål

Triatoma incrassata Usinger Figure 2 Triatoma indictiva Neiva

Triatoma lecticularia Stål

Figure 3 Triatoma neotomae Neiva

Triatoma protracta protracta Uhler

Figure 4 Triatoma protracta woodi Usinger

Triatoma recurva Stål

Figure 5 Triatoma rubida Uhler

Triatoma rubrofasciata DeGeer

Figure 6 Triatoma sanguisuga Leconte

Paratriatoma hirsuta

Adult 12-14.5 mm

Photo by Christiane Weirauch.

Triatoma gerstaeckeri

Adult 23-28.5 mm

Photo by Edward Wozniak

Figure 1. Triatoma in the United States (continued).
58 http://www.cs.amedd.army.mil/amedd_journal.aspx

12. Rassi A Jr, Rassi A, Little WC, et al. Development and validation of a risk score for predicting death in Chagas’ heart disease. N Engl J Med. 

2006;355(8):799-808.

13. Department of Defense Instruction 6130.03: Medical Standards for Appointment, Enlistment, or 

Induction in the Military Services. Washington, 

DC: US Department of Defense; April 28, 2010. 

Available at: http://dtic.mil/whs/directives/corres/

pdf/613003p.pdf. Accessed January 10, 2017.

14. Bern C, Montgomery SP, Herwaldt BL, et al. 

Evaluation and treatment of Chagas disease in 

the United States: a systematic review. JAMA. 

2007;298(18):2171-2181.

15. Wagner N, Jackson Y, Chappuis F, Posfay-Barbe 

KM. Screening and management of children at risk 

for Chagas disease in nonendemic areas. Pediatr 

Infect Dis J. 2016;35(3):335-337.

16. Sarkar S, Strutz SE, Frank DM, Rivaldi CL, Sissel B, Sánchez–Cordero V. Chagas’ disease risk in 

Texas. PLoS Negl Trop Dis. 2010;4(10):e836.

17. Garcia MN, Woc-Colburn L, Aguilar D, Hotez PJ, 

Murray KO. Historical perspectives on the epidemiology of human Chagas disease in Texas and 

recommendations for enhanced understanding 

of clinical Chagas disease in the southern United 

States. PLoS Negl Trop Dis. 2015;9(11):e0003981.

A CASE OF CHAGAS CARDIOMYOPATHY FOLLOWING INFECTION IN SOUTH CENTRAL TEXAS

Triatoma incrassata

Adult 17.5-19.8 mm

No photo available..

Triatoma indictiva

Adult 18-22 mm

Photo by Christina Wozniak.

Figure 2. Triatoma in the United States (continued).

Triatoma lecticularia

Adult 19.5-25 mm

Photo by Edward Wozniak.

Triatoma neotomae

Adult 16.5-19.5 mm

Photo by Mike Quinn

TexasEnto.net.

Figure 3. Triatoma in the United States (continued).

Triatoma protracta protracta

Adult 16-23 mm

Photo by Justin Schmidt and 

Steven Klotz.

Triatoma protracta woodi

Adult 14-16 mm

Photo by Edward Wozniak.

Figure 4. Triatoma in the United States (continued).

Figure 5. Triatoma in the United States (continued).

Triatoma recurva

Adult 25.5-33 mm

Photo by Justin Schmidt and 

Steven Klotz.

Triatoma rubida

Adult 15.5-23 mm

Photo by Edward Wozniak.
January – June 2017 59

THE ARMY MEDICAL DEPARTMENT JOURNAL

18. Klotz SA, Dorn PL, Mosbacher M, Schmidt JO. 

Kissing bugs in the United States: risk for vectorborne disease in humans. Environ Health Insights. 

2014;8(suppl 2):49-59.

19. Kjos SA, Snowden KF, Olson JK. Biogeography and Trypanosoma cruzi infection prevalence 

of Chagas disease vectors in Texas, USA. Vector 

Borne Zoonotic Dis. 2009;9(1):41-50.

20. Martínez-Ibarra JA, Alejandre-Aguilar R, Paredes-González E, et al. Biology of three species of 

North American Triatominae (Hemiptera: Reduviidae: Triatominae) fed on rabbits. Mem Inst Oswaldo Cruz. 2007;102(8):925-930.

21. Curtis-Robles R, Wozniak EJ, Auckland LD, Hamer GL, Hamer SA. Combining public health education and disease ecology research: using citizen science to assess Chagas disease entomological risk 

in Texas. PLoS Negl Trop Dis. 2015;9(12):e0004235. 

doi: 10.1371/journal.pntd.0004235.

22. Wozniak EJ, Lawrence G, Gorchakov R, et al. The 

biology of the Triatomine bugs native to south central Texas and assessment of the risk they pose for 

autochthonous Chagas disease exposure. J Parasitol. 2015;101(5):520-528.

23. Bern C, Kjos S, Yabsley MJ, Montgomery SP. Trypanosoma cruzi and Chagas disease in the United 

States. Clin Microbiol Rev. 2011;24(4):655-681.

24. Gorchakov R, Trosclair LP, Wozniak EJ, et al. Trypanosoma cruzi infection prevalence and bloodmeal analysis in Triatomine vectors of Chagas 

disease from rural peridomestic locations in Texas, 

2013-2014. J Med Entomol. 2016;53(4):911-918.

25. Shender L, Niemela M, Conrad P, Goldstein 

T, Mazet J. Habitat management to reduce human exposure to Trypanosoma cruzi and western 

conenose bugs (Triatoma protracta). Ecohealth. 

2016;13(3):525-534.

26. AFPMB Technical Guide No. 36: Personal Protective Measures Against Insects and Other Arthropods of Military Significance. Silver Spring, MD: 

Armed Forces Pest Management Board: November 

6, 2015. Available at: http://www.acq.osd.mil/eie/

afpmb/docs/techguides/tg36.pdf. Accessed January 10, 2017.

Authors

Maj Webber is the Preventive Medicine Element Chief, 

559th Trainee Health Squadron, Wilford Hall Ambulatory Surgical Center, JBSA-Lackland, Texas.

Lt Col Wozniak is Chief Public Health Officer, Texas 

State Guard Medical Brigade, Camp Mabry, Texas.

CPT Chang is a Fellow in the Department of Infectious 

Disease, San Antonio Military Medical Center, JBSAFort Sam Houston, Texas.

Maj Bush is a Fellow in the Cardiology Division, San 

Antonio Military Medical Center, JBSA-Fort Sam Houston, Texas.

Maj Wilson is Medical Director, 559th Trainee Health 

Squadron, Wilford Hall Ambulatory Surgical Center, 

JBSA-Fort Sam Houston, Texas.

LTC Watts is Chief of Cardiology, Cardiology Division, 

San Antonio Military Medical Center, JBSA-Fort Sam 

Houston, Texas.

Lt Col Yun is Infectious Disease Fellowship Program Director, San Antonio Uniformed Services Health Education Consortium, San Antonio Military Medical Center, 

JBSA-Fort Sam Houston, Texas.

Triatoma rubrofasciata

Adult 19.5-25 mm

Photo by Gabe Hamer.

Triatoma sanguisuga

Adult 15.5-23 mm

Photo by Edward Wozniak.

Figure 6. Triatoma in the United States (end).
60 http://www.cs.amedd.army.mil/amedd_journal.aspx

Leishmaniasis is recognized as a significant threat to 

deployed military forces and is a public health priority.1

It is an emerging zoonotic disease caused by obligate intracellular parasites of the genus Leishmania which are 

transmitted by Phlebotomine sand flies. Currently, over 

350 million people are at risk and 12 million people in 

90 countries are infected.2

 The number of new cases of 

the non-fatal form of the disease, cutaneous leishmaniasis (CL), and the potentially fatal visceral (VL) form 

of the disease is 1.5 and 0.5 million annually. The most 

prevalent regions at risk are the Middle East, Africa (primarily east and north), India, Asia, Central and South 

America (primarily Honduras and Brazil). Leishmaniasis is recognized as an emerging disease in the United 

States. Visceral leishmaniasis is caused by Leishmania 

donovani complex L donovani and L infantum in the Old 

World and L infantum/L chagasi in the New World. Cutaneous leishmaniasis is caused primarily by L tropica

and L major complex in the Old World and L mexicana

complex (L mexicana, L amazonensis, and L venezuelensis); and subgenus Viannia complex L brazilliensis

and L guyanensis in the New World. A licensed vaccine 

or drug prophylaxis does not exist.

Visceral leishmaniasis is fatal if untreated.3,4 Cutaneous 

leishmaniasis can cause severe scarring with permanent 

disfigurement and physical impairment.5,6 Both forms of 

the disease often require lengthy and painful treatment 

and may present complex issues associated with drug 

efficacy, adverse effects, and cost.7

 Early and accurate 

diagnosis is critical to efficacious treatment. Early treatment decreases VL mortality, CL physical impairment 

and disfigurement, and the length and expense of care.

Visceral and cutaneous leishmaniasis clinical samples 

are obtained from venous blood, spleen and bone marrow aspirate and skin lesion biopsy, respectively. Laboratory methods used for diagnoses are primarily microscopy and immunoassay.7,8 Isoenzyme analysis is 

typically used to identify Leishmania species. However, 

these methods can take days to weeks to produce results. 

Commercially available immunochromatographic (ICT) 

strip assays are available that detect L donovani antibody in human serum, providing an aid in presumptive 

diagnosis of VL. Moreover, ICT strip assays are easily 

transported and require minimal training, 2 key traits to 

providing an efficacious field-expedient methodology.9

Detection by ICT augmented by molecular-based methods achieves optimal sensitivity and specificity.10,11

Polymerase chain reaction (PCR) detection tests are 

finding a place as an aid in leishmaniasis diagnosis.12-17

However, detection by PCR is challenged by the number 

of infecting parasites from potentially low load CL patient samples of 2 to 11e6 parasites/skin biopsy and VL 

sample loads of 30 to 2e5 parasites/mL blood with a median of ≈1e3 parasites/mL.18,19 The load of CL parasites 

in blood is presumably at single log or less concentration based on PCR and isoenzyme test results.20,21

Direct Detection of Leishmania from

 Clinical Samples

John N. Waitumbi, PhD Ratree Takhampunya, PhD

MAJ Joshua Bast, MS, USA LTC Anthony L. Schuster, MS, USA

Nancy Nyakoe, MS Marshall T. Van de Wyngaerde, MS

Charles Magiri James C. McAvin, MS

Miguel Quintana, PhD Russell E. Coleman, PhD

Abstract

The ability to rapidly and accurately diagnose leishmaniasis is a military priority. Testing was conducted to 

evaluate diagnostic sensitivity and specificity of field-expedient Leishmania genus and visceral Leishmania

specific dual-fluorogenic, hydrolysis probe (TaqMan), polymerase chain reaction assays previously established 

for use in vector surveillance. Blood samples of patients with confirmed visceral leishmaniasis and controls 

without the disease from Baringo District, Kenya, were tested. Leishmania genus assay sensitivity was 100% 

(14/14) and specificity was 84% (16/19). Visceral Leishmania assay sensitivity was 93% (13/14) and specificity 

80% (4/5). Cutaneous leishmaniasis (CL) skin scrapes of patients from Honduras were also evaluated. Leishmania genus assay sensitivity was 100% (10/10). Visceral Leishmania assay specificity was 100% (10/10) from 

cutaneous leishmaniasis samples; no fluorescence above background was reported. These results show promise 

in a rapid, sensitive, and specific method for Leishmania direct detection from clinical samples.
January – June 2017 61

We describe here sensitive and specific Leishmania

genus (LEIS) and visceral Leishmania (LVL) TaqMan 

PCR assays (ThermoFisher Scientific, Waltham, MA) 

which clearly show promise as an aid in diagnosis.

Methods

Clinical Samples

Test panels were prepared consisting of well characterized nucleic acid extracts of VL confirmed patient and 

healthy control blood samples and CL confirmed patient 

skin scrapes.

Blood samples of patients with VL and controls without 

the disease were obtained in endemic areas of Baringo 

District, Kenya, during routine surveillance for VL.22

These samples were used to construct a test panel of 14 

VL patients and 19 control samples. Testing was conducted at the Basic Science Department, USAMRU-K 

in February 2012. Clearance was obtained from the 

institutional ethical committee of the Kenya Medical 

Research Institute (KEMRI ERC #1282) and the Institutional Review Board (IRB), Walter Reed Army Institute of Research (WRAIR), Silver Spring, Maryland 

(WRAIR IRB #1402).

Prior to evaluations conducted in Kenya reported here, 

LEIS and LVL assays were tested under a separate study 

using total nucleic acid extracts from isolates of 10 CL 

patient skin scrape samples obtained in Honduras. Samples were provided by the Honduras Ministry of Public 

Health. Serology was used to identify the causative agent 

as L mexicana. Extracts were prepared and transported 

to the United States Army Center for Health Promotion and Preventive Medicine-West (CHPPM-W), Fort 

Lewis, WA. Testing was conducted at the Department of 

Entomology, CHPPM-W. Test results were used for PCR 

assay evaluation purposes only. Aliquots of specimens 

routinely ordered for conventional diagnostic purposes 

were used in this study and all patient identifiers were 

removed prior to the receipt of specimens in compliance 

with the human use protocol, “Surveillance of Emerging Infectious Disease Agents in Human Populations of 

Central America,” USAMRIID, Fort Detrick, Maryland, 

approved 2001 (FY01-08).

Nucleic Acid Preparation

Extracts were prepared using commercially available, 

off the shelf, preformatted, guanidinium thiocyanate 

based total nucleic acid purification kits following 

the manufacturer’s instructions (QIAGEN, Valencia, 

California).

PCR Assay Design

Previously established Leishmania genus primer and 

probe sequences were obtained from the literature for 

LEIS freeze-dried assay development.23 Design of LVL 

primer and probe oligonucleotides and LVL and LEIS 

PCR reaction conditions were previously described.24

Leishmania genus and LVL assay optimization, limit 

of detection (LoD), analytic sensitivity, and specificity test results are previously described.24 Primer and 

probe synthesis, proprietary master mix preparation 

and freeze-drying process, and quality control were 

conducted commercially (BioFire Diagnostics Inc., Salt 

Lake City, Utah).

Analyses

The analytic platform was a Department of Defense approved, field-expedient, real-time PCR thermocycler, the 

Ruggedized Advanced Pathogen Identification Device 

(RAPID) (Idaho Technology, Inc, Salt Lake City, Utah) 

as shown in the Figure. The analytic system and PCR 

cycling conditions were previously described.24 The 

comparator test for this study was a well-established L 

donovani PCR assay targeting a 360 base-pair region of 

the 6-phosphate isomerase gene.25 Comparator test analysis was conducted using a SmartCycler (Cepheid, Inc, 

Sunnyvale, California). Blind testing was conducted.

Data Management and Statistical Analyses

Sample identification and experiment information were 

entered electronically in the RAPID operating system 

run protocol. A single data point at the end of each annealing extension cycle was collected and reported as 

TaqMan probe fluorescence released by 5’-nuclease activity during primer extension. The criterion for a positive result was a significant increase in fluorescence over 

background levels, that is, cycle threshold (Ct), defined by 

an algorithm provided in the RAPID analytical software 

(Roche Molecular Biochemicals, Basel, CH). The Ct is 

defined as the first PCR cycle with significant fluorescence when normalized against background fluorescence. 

Samples with a mean Ct more than 40 were considered 

negative, whereas samples with a mean Ct of 40 or less 

were considered positive by RAPID analyses. Sample results and statistical analyses were automatically archived.

Results

Test panels consisting of well characterized nucleic acid 

extracts of blood samples from 14 VL positive patients 

and 19 controls without VL, and 10 CL positive patient 

skin scrape samples were used to evaluate LEIS and 

LVL assay diagnostic sensitivity and specificity. The results are presented in the Table.

THE ARMY MEDICAL DEPARTMENT JOURNAL
62 http://www.cs.amedd.army.mil/amedd_journal.aspx

Visceral Leishmaniasis Diagnostic Sensitivity

from Blood Samples

Visceral leishmaniasis test results were LEIS sensitivity 100% (14/14) and specificity 84% (16/19) and LVL 

sensitivity 93% (13/14) and specificity 80% (4/5). Mean 

LEIS Ct and standard deviation were 26.96 and 4.32, respectively, where n=14, SE=1.16 (95% CI, 24.69-29.23). 

Specificity of the LEIS assay was 84% (16/19). False 

positive results occurred for 3 samples. The Ct values 

were 30.63, 30.9, and 31.48. The sensitivity of the LVL 

assay was 93% (13/14). Mean LVL Ct and standard deviation were 31.80 and 4.36, respectively, where n=13, 

SE=1.21 (95% CI, 29.43-34.17). Specificity of the LVL 

assay was 80% (4/5). One false positive occurred with a 

Ct value of 34.49.

Cutaneous Leishmaniasis Diagnostic Sensitivity

from Skin Lesion Samples

Test panel results of CL skin scrapes of patients from 

Honduras were LEIS sensitivity 100% (10/10). Honduras test panel LEIS results were mean Ct and standard 

deviation of 26.51 and 2.87, respectively, where n=10, 

SE=0.91 (95% CI, 24.73-28.29). The specificity of the 

LVL assay was 100% (10/10); no detectable fluorescence 

above background was observed.

Controls

Throughout laboratory testing and field evaluations, positive template control reactions reported fluorescence at 

the expected LEIS assay Ct value (≈18) and LVL assay 

Ct value (≈28).24 Negative template control reactions 

did not report fluorescence above background. No fluorescence above background was observed in testing of 

LEIS and LVL assay specificity using human genomic 

DNA (1ng/reaction).

Comment

This preliminary evaluation of LEIS and LVL TaqMan 

PCR assay diagnostic sensitivity and specificity shows 

promise in a rapid, sensitive, and specific method for 

Leishmania direct detection from VL patient blood and 

CL skin scrape samples. However, enhancement of our 

method and additional testing are needed.

The LEIS assay serves as a valuable screening tool 

for subsequent Leishmania species identification by 

molecular and/or isoenzyme analyses. The diagnostic 

sensitivity of the LVL assay was 93% (n=14) which is 

approximately in line with previous reports describing direct detection from VL patient blood samples 

by real-time PCR; 79% (n=15), 80% (n=45), and 94% 

(n=100).13-15 However, it is unlikely that we can reliably 

reproduce greater than 90% sensitivity results without 

enhancing our methods. False negative results are inevitable because VL patient blood samples potentially 

harbor parasites at concentrations as low as a single log. 

While PCR limit of detection is achieved at single copy 

and regularly at single log concentrations, there is no 

expectation of reliable and reproducible results from low 

load samples using conventional methods. The issue is 

that only a fraction of the sample is extracted when using commercial preformatted kits (100 μL-200 μL) or 

robotic systems (100 μL-400 μL).26-28 Thus, for samples 

with a low parasite load, the resulting extract concentration of target template is likely to be diluted below 

detection limits. Reports of ≈80% diagnostic sensitivity are likely to better represent consistently achievable 

direct detection given the limitations of conventional 

sample preparation methodologies.

In this study, the specificity of LEIS (84%, 16/19) and 

LVL (80%, 4/5) assays from VL patient blood samples 

were probably underestimated. Both LEIS and LVL 

assays reported robust fluorescence from the same 

“healthy control,” indicating incorrect reporting by the 

L donovani comparator test result. Better than reported 

LEIS assay specificity was also evidenced by subsequent 

use of our early development phase “Human Leishmania (LHL)” assay which is designed to detect VL and 

CL causative agents. Both, LEIS and LHL results were 

positive (Ct 31.48 and 35.82, respectively) and negative 

by LVL for the same healthy control, indicating that the 

sample harbored a CL causative agent.

The field-expediency and clinical implications associated with VL diagnosis from noninvasively obtained body 

fluids versus invasively obtained samples are profound. 

Direct detection of VL parasites from noninvasively 

obtained body fluids is challenged by characteristically 

single log loads. Various degrees of success have been 

achieved from urine (88%, n=17) and oral fluid (100%, 

n=37 and 83%, n=148).29-31 Detection from canine oral 

Results of LEIS and LVL RT-PCR assay sensitivity and specificity testing using clinical samples.

Disease Source Origin Sensitivity LEIS PCR

Ct Mean/SD

Specificity Sensitivity LVL PCR

Ct Mean/SD

Specificity

VL Blood Kenya 100%(14/14) 26.96/4.32 84%(16/19) 93%(13/14) 31.80/4.36 80%(4/5)

CL Lesion Honduras 100%(10/10) 26.51/2.87 100%(10/10)

VL indicates visceral leishmaniasis; CL, cutaneous leishmaniasis; Ct, critical threshold; LEIS, Leishmania genus; LVL, visceral 

Leismania; and PCR, polymerase chain reaction [detection test].

DIRECT DETECTION OF LEISHMANIA FROM CLINICAL SAMPLES
January – June 2017 63

THE ARMY MEDICAL DEPARTMENT JOURNAL

and conjunctival fluid show promise as well.32 In these 

studies, conventional sample preparation methods were 

augmented with centrifugation to concentrate parasites 

prior to extraction.

We are currently developing an alternative approach. 

Microfiltration methodology versus centrifugation mitigates variables affecting relative yields of pathogens 

obtained from patient body fluids. The specific gravity 

(SG) of body fluids varies widely across patients and for 

samples obtained from the same patient over time. It has 

been shown that centrifugation of high SG urine results 

in significantly higher yields of organism compared to 

low SG urine samples.33 To help ensure consistently high 

relative yields, we are developing a filtration approach to 

eliminate SG variables. Preliminary results of our leptospirosis diagnostic system show promise in increasing analytical limit of detection by 1-2 log concentration 

compared to centrifugation. This will be described in a 

future article. The low cost, simple, and portable characteristics of filtration technology also influenced our decision to take a microfiltration/capture approach.

The results of LEIS and LVL detection assay testing 

show promise in a rapid, sensitive, and specific method 

for Leishmania direct detection from clinical samples. 

It is our intent to augment analytical capabilities with 

standardized, field-expedient, detection enhancing, preanalytical technologies.

Acknowledgements

Our thanks to the health facilities in Kenya that provided 

Leishmania samples and the Ministry of Health of Honduras. Funding for the Kenyan study was from the Armed 

Forces Health Surveillance Branch and its Global Emerging Infections Surveillance and Response Section. This 

work was funded in part by the Military Infectious Diseases Research Program, USAMRC, Fort Detrick, Maryland, and the Air Force Medical Support Agency, Research, Development and Innovations Directorate, Office 

of the Surgeon General, Falls Church, Virginia. The opinions or assertions contained herein are the private views of 

the authors, and are not to be construed as official, or as 

reflecting true views of the Department of the Army, Department of the Navy, Department of the Air Force, or the 

Department of Defense.

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Authors

Dr Waitumbi, Ms Nyakoe, and Mr Magiri are with the 

Kenya Medical Research Institute/US Army Medical 

Research Unit, Kenya.

MAJ Bast is Deputy Director/Research Entomologist 

with the US Army Medical Research directorate - Georgia, in Tbilisi, Georgia.

Dr Quintana is with the US Army Public Health Command Region-South, Joint Base San Antonio Fort Sam 

Houston, Texas.

Dr Takhampunya is with the Armed Forces Research Institute of Medical Sciences, Bangkok, Thailand.

LTC Schuster is with the Directorate of Combat and Doctrine Development, AMEDD Center & School, Health 

Readiness Center of Excellence.

Mr Van de Wyngaerde is with the Walter Reed Institute 

of Research, Silver Spring, Maryland.

Mr McAvin, a retired US Air Force employee, is with 

Biosciences Research Management, Inc, St Louis, 

Missouri.

Dr Coleman, a retired US Army Colonel, is with the Battelle National Biodefense Institute, Frederick, Maryland.

DIRECT DETECTION OF LEISHMANIA FROM CLINICAL SAMPLES
January – June 2017 65

Phlebotomine sand flies (Subfamily Phlebotominae, 

Family Psychodidae, Order Diptera) are vicious biting 

insects that cause extreme nuisance and transmit diseases to humans. In Africa and Asia (Figure 1), the peridomestic species bite insatiably and also are of major 

public health concern in many parts of the world, particularly in the Old World, where they are capable of 

transmitting pathogens, including protozoans (Leishmania) and viruses (Phleboviruses, sand fly fever).1 Phlebotomus females suck blood, including humans, while 

Sergentomyia females primarily feed on reptiles, and 

rarely bite humans.2

 Subgenera (Adlerius, Euphlebotomus, Larroussius, Phlebotomus, Paraphlebotomus, and 

Synphlebotomus) in the genus Phlebotomus contain 

species that are vectors or suspected vectors in the Old 

World.2

 Of approximately 266 sand fly species in the Old 

World (Southwest Asia, Central Asia, and Africa including Western Indian Ocean Islands ), only about 27 species are capable of transmitting protozoan Leishmania

parasites that cause visceral leishmaniasis (kala-azar) 

and various forms of cutaneous leishmaniasis (oriental 

sore, espundia, etc) in humans.2-4 A few sand fly species have been associated with Phlebovirus and other 

viruses.3-6 Numerous reports are available on the biology, and medical importance (as disease vectors) of sand 

flies and leishmaniasis control.2,7-10 In the Old World, the 

human biting Phlebotomus sand flies and major leishmaniasis transmissions are confined in the subtropics 

(particularly in dry, semiarid areas), with some human 

biting species in Africa south of the Sahara.2

Among diseases transmitted by Phlebotomine sand flies, 

leishmaniasis has the most significant effect on military 

operations, particularly those of the United States.11 In 

Afghanistan (Operation Enduring Freedom, OEF) and 

Iraq (Operation Iraqi Freedom, OIF), numerous US Soldiers were exposed to significant leishmaniasis risk and 

about 1,287 incident diagnoses/reports of leishmaniasis, both cutaneous (1,283 cases) and visceral (4 cases) 

forms, were reported from 2001-2006 among OEF/OIF 

deployed troops.12 During OIF, US military entomologists conducted the sand fly surveillance at Tallil Air 

Base, Iraq, from April 2003 to November 2004, and they 

noted the strong effect of sand fly vectors on military 

operations, including the leishmanial threat to deployed 

troops in Iraq.13-15

In this article, we examine the types and related specimens of Old World Phlebotomine sand flies deposited in 

the following: US National Museum of Natural History 

New Records, Distribution, and Updated

 Checklists of Old World Phlebotomine

 Sand Flies, With Emphasis on Africa,

 Southwest Asia, and Central Asia

Leopoldo M. Rueda, PhD Sarah Benyamin, BS

James E. Pecor, BS Philippe Boussès, PhD

Matthew Wolkoff, BA Mustapha Debboun, PhD, BCE

David Pecor, BS

Abstract

This article includes new records, distribution, and updated checklist of Phlebotomine sand flies (Psychodidae, 

Diptera) in the Old World (Africa including West Indian Ocean Islands, Southwest Asia, and Central Asia) 

based on specimen collections housed in different repositories worldwide. About 124 species have primary 

types housed in 5 repositories including holotypes (45 species, 4 subspecies), syntypes (28 species, 3 subspecies), “types” (14 species), allotypes (10 species), paratypes (36 species, 3 subspecies), lectotypes (13 species), 

and cotype (5 species), mounted on 671 slides. New abbreviations were proposed for 2 subgenera in the genus 

Phlebotomus and 6 subgenera in the genus Sergentomyia. New country records were noted in Phlebotomus (4 

species in 4 subgenera in 7 countries) and Sergentomyia (10 species in 4 subgenera in 8 countries). For species 

diversity in the Old World, Phlebotomus includes 92 species and 7 subspecies in 9 subgenera, while Sergentomyia includes 166 species and 16 subspecies in 12 subgenera. A total of 95 species and 7 subspecies of 2 

genera (Phlebotomus and Sergentomyia) were recorded in Africa while about 26 species and 16 subspecies in 

Southwest Asia and Central Asia.
66 http://www.cs.amedd.army.mil/amedd_journal.aspx

(USNMNH), Smithsonian Institution, Suitland, MD; 

The Museum of Natural History (MNH), London, UK; 

Institut de Recherche pour le Développement (IRD), 

Montpellier, France; Musée National d’Histoire Naturelle Entomologie (MNHE), Paris, France; Institut 

Pasteur (IP), Paris, France; Musée Royal de L’Afrique 

Centrale (MRAC), Dept Entomologie, Tervuren, Belgium. Additional specimens were borrowed from the 

following institutions: Reimer University (RU), Reimer, France; US Army/Kenya Medical Research Institute 

(KEMRI), Nairobi, Kenya; Florida State Collection of 

Arthropods (FSCA), Gainesville, Florida; and US Navy 

laboratories (UNL). In addition, 8 interactive computerized identification keys (LUCID software16) of selected 

African and Asian sand fly species (potential or known 

vectors), based on specimens from different depositories 

are presently posted on the Walter Reed Biosystematics 

Unit (WRBU) website (wrbu.si.edu).17

We record the collection data of sand flies, including 

their geographical distribution, past and present taxonomic arrangement, and related information. The species occurrence and diversity of these sand flies, according to the number of collections for each country 

over certain periods, were analyzed and reported. Other 

collections or occurrence data of sand fly specimens (including non-types, from the Old World) from the 6 repositories (USNMNH, MNH, IRD, MNHE, IP, MRC) 

were also examined and recorded, and will be posted 

to the WRBU/VectorMap website (vectormap.si.edu).18 These data may be helpful in 

developing world sand fly taxonomic catalogs, sand fly vector risk maps, and predictive distribution models. In addition to 

increasing the knowledge of sand fly distribution, the collection holdings in these repositories, particularly the primary types, 

will assist future phlebotomine researchers 

in their taxonomic and related studies.

Materials and Methods

Old World Phlebotomine sand fly specimens used in this study were deposited 

in 6 repositories (USNMNH, MNH, IRD, 

MNHE, IP, MRC), or borrowed from various agencies (RU, KEMRI, FSCA,UNL). 

The slide mounted specimens were examined and their collection data were recorded. All collection data from selected 

repositories were recorded verbatim and 

entered into a standard specimen label data 

capture spreadsheet. The data were georeferenced using the point-radius method of 

georeferencing locality descriptions that 

do not have associated geographic coordinates. The 

georeferenced data were processed and used for analyses. The primary types (holotypes, allotypes, paratypes, 

syntypes) of sand flies were examined for collection records and related information. Photographs of primary 

types of selected sand fly species were taken, and will 

be processed for another report.

Results

Species Types and Abbreviations

The proposed abbreviations for genera and subgenera 

of Phlebotominae (Psychodidae, Diptera) of the Old 

World is shown in Table 1, based primarily on Marcondes,20 with additions of new abbreviations for the 2 

subgenera in genus Phlebotomus, ie, Legeromyia, Leg.; 

Madaphlebotomus, Mad.; and the 6 subgenera in genus 

Sergentomyia, ie, Capensomyia, Cap.; Parvidens, Pav.; 

Rondanomyia, Ron.; Spelaeomyia, Spe.; Trouilletomyia, 

Tro., and Vattieromyia, Vat.

The list of Old World Phlebotomine species with type 

specimens deposited in the various repositories is 

shown in Table 2 (see pages 71-73), using the available 

taxonomic arrangements.19 The number of slides for 

each type, species, sex, and the country of type origin 

are also included in Table 2. About 124 species have 

primary types deposited in 5 repositories (MNH, IRD, 

MNHE, IP, MRC), including holotypes (45 species, 4 

subspecies), syntypes (28 species, 3 subspecies), “types” 

NEW RECORDS, DISTRIBUTION, AND UPDATED CHECKLISTS OF OLD WORLD

PHLEBOTOMINE SAND FLIES, WITH EMPHASIS ON AFRICA, SOUTHWEST ASIA, AND CENTRAL ASIA

Figure 1. Map showing different countries in the Old World, particularly Africa, 

Southwest Asia, and Central Asia, and respective country codes (see Table 4 for 

code–country definitions). Note: a star indicates the location of various primary 

types.
January – June 2017 67

THE ARMY MEDICAL DEPARTMENT JOURNAL

(14 species), allotypes (10 species), paratypes (36 species, 3 subspecies), lectotypes (13 species), and cotype 

(5 species), mounted on 671 slides. Those specimens on 

slides labeled “Type” could be considered as holotypes, 

but proper designations should be done later.

New Country Records and Updated Checklist of 

Phlebotomine Species

An updated checklist of Phlebotomine sand fly species 

in different countries of Africa (including Western Indian Ocean Islands), Southwest Asia, and Central Asia 

is presented in Table 3 (see pages 74-82). It also includes 

distribution of these species in Africa (58 countries), 

Southwest and Central Asia (26 countries) and other 

countries (35), as well as their selected references.3,9,32-49

Several islands of Western Indian Ocean, ie, Comoros, 

Madagascar, Mauritius, Mayotte (France), Réunion 

(France), and Seychelles are included as Africa for the 

purposes of this article.50

The genus Phlebotomus includes 

92 species and 7 subspecies in 

9 subgenera (Adlerius, 17 species; Anaphlebotomus, 6 species; 

Euphlebotomus, 5 species; Larroussius, 27 species, 5 subspecies); Legeromyia, 1 species; 

Madaphlebotomus, 4 species; 

Paraphlebotomus, 12 species, 2 

subspecies; Phlebotomus, 6 species; Synphlebotomus, 10 species; 

Transphlebotomus, 2 species). 

The genus Sergentomyia includes 

166 species and 16 subspecies in 

12 subgenera (Capensomyia, 10 

species; Demeillonius, 1 species; 

Grassomyia, 6 species, 2 subspecies; Neophlebotomus, 22 species; 

Parrotomyia, 16 species, 4 subspecies; Rondanomyia, 1 species, 

2 subspecies; Sergentomyia, 45 

species, 6 subspecies; Sintonius, 

27 species; Spelaeomyia, 6 species; Trouilletomyia, 2 species; 

Vattieromyia, 4 species), and one 

ungrouped taxon (24 species). 

Further, we recorded a total of 

95 species and 7 subspecies of 2 

genera (Phlebotomus and Sergentomyia) in Africa, while about 26 

species and 16 subspecies in Southwest Asia and Central 

Asia. About 22 species of both genera had wide distributions and were found in 35 countries other than Africa, 

Southwest Asia, and Central Asia.

The new records of species in different countries are 

shown in Table 3 (with country codes (CN) as listed in 

Table 4, see page 83). In the genus Phlebotomus, 4 species in 4 subgenera were recorded for the first time in 7 

countries. These species included the following: Phlebotomus (Adlerius) chinensis Newstead, Iran (CN 66); 

Ph. (Anaphlebotomus) rodhaini Parrot, Côte d’Ivoire 

(CN 14), Ghana (CN 22), Mozambique (CN 36); Ph. 

(Paraphlebotomus) jacusieli Theodor, Lebanon (CN 

73); and Ph. (Phl.) papatasi (Scopoli), Kenya (CN 25). In 

the genus Sergentomyia, 10 species in 4 subgenera were 

recorded for the first time in 8 countries. These species 

included the following: Sergentomyia (Grassomyia) 

ghesquierei (Parrot), Ghana (CN 22) Somalia (CN 47); 

Se. (Neophlebotomus) decipiens (Theodor), Tanzania 

(CN 52); Se. (Neo.) dureni Parrot, Ethiopia (CN 19); Se. 

(Par.) freetownensis freetownensis (Sinton), Democratic 

Republic of Congo (CN 12); Se. (Par.) magna (Sinton) 

Ghana (CN 22); Se. (Sergentomyia) antennata (Newstead), Tanzania (CN 52); Se. (Ser.) buxtoni

(Theodor), Côte d’Ivoire CN (14); 

Se. (Ser.) magnidentata Davidson, 

Kenya (CN 25); Se. (Ser.) minuta

minuta Rondani, Kenya (CN 25); 

and Se. (Ser.) schwetzi (Adler, 

Theodor & Parrot), Tanzania (CN 

52).

Three taxa46 were not included in 

the updated list shown in Table 

3 pending clarification of their 

taxonomic identities, namely: Se. 

(Ser.) bereiri, Se. (Ser.) distinctus (as distincta), and Se. (Ser.) 

fermatus (as firmata). They cannot be treated as subspecies of Se. 

(Ser.) bedfordi because of their 

overlapping distributions.3 These 

3 taxa were treated by Davidson51

as good species based on a unique 

and maybe aberrant holotype female of Se. bedfordi. Seccombe et 

al3

 did not support it due to the 

absence of statistical and/or biological analysis of sympatric populations. They noted that the morphological, ecological, and geographical data of Davidson51 may 

help in future taxonomic studies. 

Niang et al 46 provided diagnostic morphological features and distribution records of these 3 taxa, in addition 

to Se. bedfordi. Although it was suggested46,51 that the 

distribution of Se. bedfordi is confined to South Africa, 

Table 1. Proposed abbreviations for genera and 

subgenera of Phlebotominae (Psychodidae, Diptera) of the Old World.

Genus Subgenus Abbreviation*

Phlebotomus Ph.

Adlerius Adl.

Anaphlebotomus Ana.

Euphlebotomus Euo.

Larroussius Lar.

Legeromyia Leg.**

Madaphlebotomus Mad.**

Paraphlebotomus Par.

Phlebotomus Phl.

Synphlebotomus Syn.

Transphlebotomus Tra.

Sergentomyia Se.

Capensomyia Cap.**

Grassomyia Gra.

Neophelebotomus Neo.

Parrotomyia Par.

Parvidens Pav.**

Rondanomyia Ron.**

Sergentomyia Ser.

Sintonius Sin.

Spelaeomyia Spe.**

Trouilletomyia Tro.**

Vattieromyia Vat.**

Chinius Ch.

*Primarily based on Marcondes.29

**Proposed new abbreviation
68 http://www.cs.amedd.army.mil/amedd_journal.aspx

other reports3,19,47 and recent examinations of specimens from different depositories or museums showed 

its wider distribution (Table 3). Recent analysis49 of the 

molecular taxonomy of Phlebotomine sand flies based 

on 3 mitochondrial gene fragments (mtDNA) confirmed 

the presence of Se. congolensis (Bequart & Walravens), 

Se. salisburiensis (Abonnenc) and Se. bedfordi “Maun” 

form from Botswana. Kruger 49 proposed this form for 

Se. caliginosa Davidson and unassigned specimens of 

Se. bedfordi group found in Botswana. Since no formal 

name is available for this form, we temporarily include 

it with Se. bedfordi in Table 3. Unless the taxonomy 

of this group is well clarified, routine identification of 

these taxa will be very difficult and their distribution 

records will not be accurate.

Comment

The Phlebotomine sand fly collections, including primary types and voucher specimens, are very important 

for vector identifications, surveillance, and control efforts. Different repositories (USNMNH, MNH, IRD, 

MNHE, IP, MRC) have voucher specimens of the major species incriminated and/or suspected as vectors7

 of 

various Leishmania species involved in the transmission 

of human leishmaniasis in the Old World. Nine of 15 

suspected vector species and 6 of 10 incriminated vector species9

 of Leishmania in the Old World have primary types deposited in the 2 repositories (MNH, IP). 

For example, MNH has the holotypes and/or “syntypes” 

of incriminated vectors, ie, Phlebotomus (Larroussius) 

perniciosus Newstead, Ph. (Lar.) orientalis Parrot, Ph. 

(Phlebotomus) duboscqi Neveu-Lemaire, and suspected vectors, ie, Ph. (Synphlebotomus) celiae Minter, Ph. 

(Lar.) perfiliewi perfiliewi Parrot, Ph. (Lar.) smirnovi

Perfil’ev, Ph. (Lar.) tobbi Adler & Theodor). Considering the importance of the distribution records of sand 

flies, particularly vector species, there is an exigent need 

to conduct additional collections of sand flies from the 

Old World in order to obtain fresh voucher specimens 

for both molecular and morphological studies, and for 

safe deposits in the USNMNH and other pertinent 

repositories.

Concerning specimen collections from various repositories, they were used in the development of LUCID interactive keys for the Old World Phlebotomine sand flies,24

particularly Africa, Southwest Asia, and Central Asia 

(US Africa Command (AFRICOM), Central Command 

(CENTCOM), Europe Command (EUCOM)). Eight morphological keys for males and females of the Old World 

were created by author L. M. Rueda, with assistance 

from WRBU staff (particularly J. Stoffer for Automontage images), and are posted at the WRBU website17 as:

3⁄4 Key to identification of male and female adult Phlebotomine sand fly genera of the Old World (PACOM [Pacific Command], CENTCOM, EUCOM, 

AFRICOM).

3⁄4 Key to identification of male and female adult Phlebotomus subgenera of Southwest and Central Asia 

(PACOM, CENTCOM, EUCOM, AFRICOM).

3⁄4 Key to identification of male adult species of subgenus Larroussius of Phlebotomus, Africa (AFRICOM), with emphasis on medically important 

species.

3⁄4 Key to identification of male adult species of subgenus Paraphlebotomus of Phlebotomus, Africa 

(AFRICOM), with emphasis on medically important species.

3⁄4 Key to identification of male adult species of subgenus Phlebotomus of Phlebotomus, Africa (AFRICOM), with emphasis on medically important 

species.

3⁄4 Key to identification of male adult species of subgenus Larroussius of Phlebotomus, Southwest and 

Central Asia (CENTCOM), with emphasis on medically important species.

3⁄4 Key to identification of male adult species of subgenus Paraphlebotomus of Phlebotomus, Southwest 

and Central Asia (CENTCOM), with emphasis on 

medically important species.

3⁄4 Key to identification of male adult species of subgenus Phlebotomus of Phlebotomus, Southwest 

and Central Asia (CENTCOM), with emphasis on 

medically important species.

An example of a screen shot of LUCID interactive key 

for species of Phlebotomus (Larroussius) male adults 

from Africa is shown in Figure 2, and posted at the 

WRBU website.17 Male genitalia are important morphological characters to identify different species of sand 

flies. Examples are shown in Figures 3 and 4 for primary 

types of Phlebotomine sand flies, ie, Ph. (Paraphlebotomus) alexandri Sinton, paratype; Ph. (Adlerius) angustus Artemiev, paratype; Ph. (Larroussius) aculeatus

Lewis, Minter & Ashford, paratype; Ph. (Lar.) longipes

Parrot & Martin, paratype; Ph. (Phlebotomus) minteri

Lewis, holotype; Ph. (Synphlebotomus) celiae Minter, 

holotype; Ph. (Syn.) vansomerenae Heisch, Guggisberg 

& Teesdale, paratype; and Se. (Sergentomyia) ashfordi

Davidson, paratype.

About 12,000 slides of Phlebotomine sand flies were donated by COL (Ret) Philip Lawyer to the USNMNH for 

safe keeping. These slides were temporarily mounted 

NEW RECORDS, DISTRIBUTION, AND UPDATED CHECKLISTS OF OLD WORLD

PHLEBOTOMINE SAND FLIES, WITH EMPHASIS ON AFRICA, SOUTHWEST ASIA, AND CENTRAL ASIA
January – June 2017 69

THE ARMY MEDICAL DEPARTMENT JOURNAL

using Hoyer’s medium before, and they will be remounted permanently and identified correctly. Their locality 

data from slide labels and collection sheets will be retrieved, recorded, and processed. Additional collection 

data from Old World countries generated by ongoing US 

military entomological surveillance efforts and newly 

published scientific literature will be entered into VectorMap regularly, to enable further analysis of species 

diversity and to create sand fly vector distribution models that will be useful for leishmaniasis risk assessments.

Sand fly slides in various repositories or museums labeled as “types” should be carefully examined, and if 

necessary, must be designated as holotype or appropriate type to avoid future confusions in taxonomic clarification and related systematic studies. For instance, the 

taxa/species without designated “holotype,” but which 

have slides labeled “syntypes” or “types” should have 

proper designations.

Acknowledgments

We express our sincere appreciation and gratitude 

to the following individuals for the loans of sand fly 

types and other specimens, and for their hospitality 

during the visits of Dr Rueda to examine their collections:

D. Fontenille, F. Simard and N. Rahola, Maladies Infectieuses et Vecteurs: Ecologie, Génétique, Evolution 

et Contrôle, Institut de Recherche pour le Développement, Montpellier, France; D. Sivell, T. M. Howard and 

Z. Adams, Entomological Collections, Molecular Collections and Plants, Department of Life Sciences, The Natural History Museum, London, UK; E. Coninck and H. 

Stephane, Musée Royal de L’Afrique Centrale, Dept. Entomologie, Tervuren, Belgium; C. Daugeron, Musée National d’Histoire Naturelle Entomologie, Paris, France; 

P. Reiter, A.-B. Failloux, J. C. Ganthier, Institut Pasteur, 

Paris, France. We also thank K.A. Barbara, J.W. Diclaro, 

II, and P.J. Obenauer, US Navy Laboratories; N. Leger, 

Reimer University, Reimer, France; and J. Bast and R. 

Sang, US Army Kenya Medical Research Institute, Nairobi, Kenya, for sand fly slides loaned and deposited at 

the US National Museum of Natural History, Washington, DC.

This research was performed under a Memorandum of 

Agreement between the Walter Reed Army Institute of 

Research and the Smithsonian Institution, with institutional support provided by both organizations. The opinions contained herein are those of the authors and do not 

reflect official views of the supporting agencies.

Figure 2. Example of screen shot of LUCID16 interactive key for species of Phlebotomus (Larroussius) male adults, Africa (source: 

US Africa Command).
70 http://www.cs.amedd.army.mil/amedd_journal.aspx

NEW RECORDS, DISTRIBUTION, AND UPDATED CHECKLISTS OF OLD WORLD

PHLEBOTOMINE SAND FLIES, WITH EMPHASIS ON AFRICA, SOUTHWEST ASIA, AND CENTRAL ASIA

Figure 3. Male genitalia of Phlebotomus sand flies. (A). Phlebotomus (Paraphlebotomus) alexandri Sinton, paratype; (B). Phlebotomus (Adlerius) angustus Artemiev, paratype; (C). Phlebotomus (Larroussius) 

aculeatus Lewis, Minter & Ashford, paratype; (D). Phlebotomus (Phlebotomus) minteri Lewis, holotype.

Figure 4. Male genitalia of Phlebotomus and Sergentomyia sand flies. (A). Phlebotomus (Larroussius) 

longipes Parrot & Martin, paratype; (B). Phlebotomus (Synphlebotomus) celiae Minter, holotype; (C). 

Sergentomyia (Sergentomyia) ashfordi Davidson, paratype; (D). Phlebotomus (Synphlebotomus) vansomerenae Heisch, Guggisberg & Teesdale, paratype.
January – June 2017 71

THE ARMY MEDICAL DEPARTMENT JOURNAL

Table 2 (part 1 of 3). Types of sand flies (Phlebotominae, Psychodidae) from Africa and Southwest and Central 

Asia, examined from different repositories, including country of type origin.

Species aDepository bType (number of slides by csex)

Country of Type Origin

Phlebotomus aculeatus NH: P (1 F, 1 M) Kenya

Phlebotomus alexandri NH: P (3 F) Iraq

Phlebotomus angustus NH: P (1 F, 5 M, 4 U) Afghanistan

Phlebotomus arabicus NH: L (M); PL (5 F) Yemen

Phlebotomus ariasi NH: P (14 M) Ethiopia

Phlebotomus ashfordi NH: H (F); P (1 F, 1 M) Ethiopia

Phlebotomus bergeroti NH: S (2 F, 4 M) Algeria

Phlebotomus caudatus NH: H (M) Afghanistan

Phlebotomus celiae NH: H (M) Kenya

Phlebotomus chabaudi IP: H (M); IR: P (2 U) Tunisia

Phlebotomus chadlii NH: P (3 F, 4 M) Kenya

Phlebotomus comatus NH: P (2 M) Afghanistan

Phlebotomus duboscqi NH: H (M) Mauritania

Phlebotomus elgonensis ME: P (1 F, 1 M) Kenya

Phlebotomus fantalensis NH: H (M), P (6 M) Ethiopia

Phlebotomus gibiensis NH: H (M); P (6 F, 20 M) Ethiopia (H; P: 6 F, 16 M); Kenya (4 M)

Phlebotomus gigas BE: L (F); T (1 F, 1M) Dem. Rep. Congo

Phlebotomus guggisbergi NH: L (M); PL (3 F, 1 M) Kenya

Phlebotomus halepensis NH: S (5 M) Iran

Phlebotomus jacusieli NH: H (M) Israel

Phlebotomus kabulensis NH: P (2 M) Afghanistan

Phlebotomus kandelakii burneyi NH: H (M); P (11 F, 23 M) Pakistan

Phlebotomus katangensis BE: T (1 M) Dem. Rep. Congo

Phlebotomus kyreniae NH: S (11 F, 9 M) Cyprus

Phlebotomus longipes NH: S (3 F, 6 M) Ethiopia

Phlebotomus marismortui NH: S (4 F, 1 M) Israel

Phlebotomus mesghalii NH: H (M); P (15 F, 12 M) Kenya (H; P: 13 F, 8 M); Iran (P: 2 F, 4 M)

Phlebotomus minteri NH: H (M); P (1 M) Tanzania

Phlebotomus mongolensis NH: S (1 F, 1 M) Afghanistan

Phlebotomus naqbenius NH: H (M), P (10 F, 10 M) Saudi Arabia

Phlebotomus nuri NH: H (M); P (17 M) Pakistan

Phlebotomus orientalis NH: S (1 F, 2 M) Ethiopia

Phlebotomus pedifer NH: H (M); P (4 F, 4 M) Kenya

Phlebotomus perfiliewi galilaeus NH: S (10 F, 13 M) Cyprus (S: 1 F, 5 M); Israel (S: 9 F, 8 M)

Phlebotomus perfiliewi perfiliewi NH: S (2 M) Ukraine

Phlebotomus perniciosus NH: S (6 M) Malta

Phlebotomus rodhaini BE: T (1 M); A (F) Mozambique

Phlebotomus rupester NH: P (1 F, 4 M) Afghanistan

Phlebotomus saevus NH: S (1 F) Ethiopia

Phlebotomus salangensis NH: P (1 F, 10 M) Afghanistan

Phlebotomus smirnovi NH: S (1 M) Belarus

Phlebotomus syriacus NH: S (5 F, 2 M) Israel

Phlebotomus tobbi NH: S (10 F, 16 M, 11 U) Iran (S: 5 F, 5 M); Israel (S: 5 F, 11 M, 9 U)

Phlebotomus vansomerenae NH: P (1 M) Kenya

Phlebotomus wenyoni NH:: S (3 M) Iran

aDepository Codes:

BE - Royal Museum of Central Africa, Tervuren, Belgium ME - Museum of Entomology, Florida State Collection of

IP - Institut Pasteur, Paris, France Arthropods

IR - Inst. Res. Dev., Montpellier, France NH - Museum of Natural History, London, UK

bType Codes:

H - holotype (1 specimen) PL - paralectotype

L - lectotype (1 specimen) S - syntype

P - paratype T - type

cSex Codes:

F - female

M - male

U - undetermined
72 http://www.cs.amedd.army.mil/amedd_journal.aspx

NEW RECORDS, DISTRIBUTION, AND UPDATED CHECKLISTS OF OLD WORLD

PHLEBOTOMINE SAND FLIES, WITH EMPHASIS ON AFRICA, SOUTHWEST ASIA, AND CENTRAL ASIA

Table 2 (continued; part 2 of 3). Types of sand flies (Phlebotominae, Psychodidae) from Africa and Southwest and Central Asia, 

examined from different repositories, including country of type origin.

Species aDepository bType (number of slides by csex)

Country of Type Origin

Sergentomyia adleri NH: S (1 M) Ghana

Sergentomyia africana africana NH: L (1 F); P (1 F) Nigeria

Sergentomyia africana asiatica NH: S (4 F, 7 M) Palestine

Sergentomyia ansarii IR: T (1 M, 1F) Iran

Sergentomyia antennata NH: H (F); P (1 M) Ghana

Sergentomyia azizi NH: S (1 F) Cyprus

Sergentomyia baghdadis NH: S (12 F, 10 M) Iraq

Sergentomyia balmicola NH: P (1 M) Cameroon

Sergentomyia bedfordi NH: H (F); BE: P (7 F, 18 M) South Africa (H: F); D. R. Congo (P: 7 F, 18 M)

Sergentomyia blossi NH: S (2 F) Kenya

Sergentomyia buxtoni NH: L (F); P (2 M); P (2 M) Ghana (L: f), P: 2 M; Mali (P: 2 M)

Sergentomyia christophersi NH: L (F); P (1 F) Pakistan

Sergentomyia clastrieri NH: P (1 F, 1 M) Guinea

Sergentomyia clydei NH: L (F); P (1 M) Pakistan

Sergentomyia collarti BE: C (1 F, 1 M); NH: S (6 F, 5 M) Dem. Rep. Congo

Sergentomyia corneti IR: H (F); P (1 F) Senegal

Sergentomyia crosarai BE: C (7 F) Dem. Rep. Congo

Sergentomyia darlingi IP: P (1 F, 1 M); IR: P (4 U) Sudan

Sergentomyia decipiens BE:T (11 F, 13 M) Dem. Rep. Congo

Sergentomyia dentata dentata NH: H (M); P (2 F) Pakistan

Sergentomyia diapagai IR: P (1 M); NH: P (1 M) Burkina Faso

Sergentomyia dolichopa IP: H (M); A (F); P (1 F, 1 M) Djibouti

Sergentomyia dubia NH: S (3 F) Ghana (S: 2 F); Nigeria (1 F)

Sergentomyia dureni BE: T (1 M); NH: S (1 F) Dem. Rep. Congo

Sergentomyia dyemkoumai IR: T (M) Côte d'Ivoire

Sergentomyia edentulus IR: H (F); P (2 F) Senegal

Sergentomyia emilii IR: H (M); P (4 M); IP: P (1 M) Rep. Congo (H: M, P: 4 M); Dem. Rep. Congo (P: 1 M)

Sergentomyia fallax cypriotica NH: S (1 F, 1 M) Cyprus

Sergentomyia formica NH: P (1 F) Namibia

Sergentomyia freetownensis freetownensis NH: H (F) Sierra Leone

Sergentomyia freetownensis furanus NH: H (F) Sudan

Sergentomyia garnhami NH: P (2 M) Kenya

Sergentomyia gracilis NH: L (1 F); P (1 M) Kenya

Sergentomyia graingeri NH: H (F) Kenya

Sergentomyia grilloti IR: H (F) Rep. Congo

Sergentomyia grjebinei IR: H (F); A (M) Rep. Congo

Sergentomyia hamoni IP: H (M); A (F) Senegal

Sergentomyia harveyi NH; H (F); P (6 F, 2 M) Kenya

Sergentomyia heischi NH: H (F) Kenya

Sergentomyia herollandi NH: P (F) Mali

Sergentomyia hodgsoni NH: L (F); P (1 M) Pakistan

Sergentomyia inermis IR: A (M); NH: S (11 F) Guinea (M); Nigeria (11 F)

aDepository Codes:

BE - Royal Museum of Central Africa, Tervuren, Belgium ME - Museum of Entomology, Florida State Collection of Arthropods

IP - Institut Pasteur, Paris, France NH - Museum of Natural History, London, UK

IR - Inst. Res. Dev., Montpellier, France NM - National Museum of Natural History, Paris

KE - Kenya National Museum, Nairobi UM - US National Museum of Natural History, Smithsonian Institution

 Suitland, Maryland

bType Codes:

A - allotype (1 specimen) P - paratype

C - cotype PL - paralectotype

H - holotype (1 specimen) S - syntype

L - lectotype (1 specimen) T - type

cSex Codes:

F - female

M - male

U - undetermined
January – June 2017 73

THE ARMY MEDICAL DEPARTMENT JOURNAL

Table 2 (continued; part 3 of 3). Types of sand flies (Phlebotominae, Psychodidae) from Africa and Southwest and 

Central Asia, examined from different repositories, including country of type origin.

Species aDepository bType (number of slides by csex)

Country of Type Origin

Sergentomyia ingrami NH: H (M); P (1 M) Ghana

Sergentomyia iranica NH: H (F); P (2 M) Iran

Sergentomyia kirki NH: S (2 F) South Sudan

Sergentomyia kitonyii NH: H (F); P (8 F) Kenya

Sergentomyia lesleyae NH: H (F); P (1 M) Sudan

Sergentomyia lewisiana IR: H (F); A (M); P (5 F, 8 M) Senegal 

Sergentomyia logonensis IR: T (F) Cameroon

Sergentomyia machadoi IR: P (1 M); NH: P (5 F, 2 M) Angola

Sergentomyia macinthosi IR: H (F) South Africa

Sergentomyia madagascariensis IR: H (M); A (F); P (1 F); NH: P (2 F) Madagascar

Sergentomyia magnidentata NH: H (F); P (1 F) Kenya

Sergentomyia meilloni NH: H (M) South Africa

Sergentomyia minuta NH: P (2 F, 5 M) Ghana (1 M, 1 F); Zimbabwe (1 F, 4 M)

Sergentomyia mirabilis BE: T (1 F, 1 M) Dem. Rep. Congo

Sergentomyia montana NH: H (F); P (1 F) Ethiopia

Sergentomyia moucheti IR: H (F) Cameroon

Sergentomyia ovazzai IR: H (F) Guinea

Sergentomyia palestinensis NH: H (F) Palestine

Sergentomyia punjabensis NH: S (1 M) Pakistan

Sergentomyia renauxi BE: P (1 F) Dem. Rep. Congo

Sergentomyia richardi BE: H (1 F) Dem. Rep. Congo

Sergentomyia rimi NH: P (2 F, 1 M) Namibia (1 F, 1M); South Africa (1 F)

Sergentomyia roberti IR: P (F) Dem. Rep. Congo

Sergentomyia rogeri IR: H (F); A (M); P (3 F) Senegal 

Sergentomyia rosannae NH: H (F); P (3 F, 4 M) Kenya

Sergentomyia schoutedeni BE: L (M); P (3 M); C (2 F) Dem. Rep. Congo

Sergentomyia schwetzi BE: C (5 F, 1 M); NH: S (1 M) Dem. Rep. Congo

Sergentomyia sidioliensis IR: H (F); A (M); P (4 M) Senegal 

Sergentomyia simillima NM: P (2 M) Ivory Coast

Sergentomyia sonyae NH: H (F); P (2 F, 5 M) Oman

Sergentomyia squamipleuris NH: L (1 F) Sudan

Sergentomyia taizi NH: H (F); P (2 F) Yemen

Sergentomyia tauffliebi IP: H (M); A (F); P (1 M) Senegal

Sergentomyia teesdalei NM: T (1 M) Kenya

Sergentomyia thomsoni NH: P (1 F) Malawi

Sergentomyia tiberiadis NH: L (F) Israel

Sergentomyia trouilleti IR: T (F) Rep. Congo

Sergentomyia wansoni BE: T (1 F, 1 M); C (1 F, 1 M) Dem. Rep. Congo

Sergentomyia yusafi NH: L (M); P (1 F) Kenya

Sergentomyia yvonnae BE: T (F) Dem. Rep. Congo

aDepository Codes:

BE - Royal Museum of Central Africa, Tervuren, Belgium ME - Museum of Entomology, Florida State Collection of Arthropods

IP - Institut Pasteur, Paris, France NH - Museum of Natural History, London, UK

IR - Inst. Res. Dev., Montpellier, France NM - National Museum of Natural History, Paris

KE - Kenya National Museum, Nairobi UM - US National Museum of Natural History, Smithsonian Institution

 Suitland, Maryland

bType Codes:

A - allotype (1 specimen) P - paratype

C - cotype PL - paralectotype

H - holotype (1 specimen) S - syntype

L - lectotype (1 specimen) T - type

cSex Codes:

F - female

M - male

U - undetermined
74 http://www.cs.amedd.army.mil/amedd_journal.aspx

NEW RECORDS, DISTRIBUTION, AND UPDATED CHECKLISTS OF OLD WORLD

PHLEBOTOMINE SAND FLIES, WITH EMPHASIS ON AFRICA, SOUTHWEST ASIA, AND CENTRAL ASIA

Table 3 (part 1 of 9). Checklist of Phlebotomine sand flies of Africa, Southwest and Central Asia, including distribution. Note: 

countries are represented by numerical codes which are defined in Table 4.

Species Africa* Southwest

Asia and

Central Asia*

Other

Countries

Selected

References

Phlebotomus (Adlerius) angustus Artemiev, 1978 60, 77, 81, 84 3, 19

Phlebotomus (Adlerius) arabicus Theodor, 1953 10, 19, 27 64,77, 85 3, 19, 46, 47, X

Phlebotomus (Adlerius) balcanicus Theodor, 1958 62, 65, 66, 80 96, 102, 109, 

111, 118

3, 19, 43, 47

Phlebotomus (Adlerius) brevis Theodor & Mesghali, 1964 62, 65, 66, 80 96, 104 3, 19, 43, 45

Phlebotomus (Adlerius) chinensis Newstead, 1916 66* X

Phlebotomus (Adlerius) comatus Artemiev, 1978 60, 75 107 3, 19

Phlebotomus (Adlerius) halepensis Theodor, 1958 62, 65, 68, 66, 

78, 80

107 3, 19, 43

Phlebotomus (Adlerius) hindustanicus Theodor, 1958 65, 75 98, 107 3, 19

Phlebotomus (Adlerius) kabulensis Artemiev, 1978 60, 65, 66 3, 19, 3

Phlebotomus (Adlerius) kyreniae Theodor, 1958 80 94 3, 19

Phlebotomus (Adlerius) longiductus Parrot, 1928 60, 66, 75, 84 92, 98, 109 3, 19, 43

Phlebotomus (Adlerius) naqbenius Lewis & Buttiker, 1986 77 3, 19

Phlebotomus (Adlerius) rupester Artemiev, 1978 60 3, 19

Phlebotomus (Adlerius) salangensis Artemiev, 1978 60, 66 3, 19

Phlebotomus (Adlerius) simici Nitzulescu, 1931 66, 68, 78, 80 96, 102 3, 19

Phlebotomus (Adlerius) turanicus Artemiev, 1974 60, 79, 81, 84 3, 19

Phlebotomus (Adlerius) zulfagarensis Artemiev, 1978 66, 81 3, 19

Phlebotomus (Anaphlebotomus) colabaensis Young & Chalam, 1927 75 98 3, 19

Phlebotomus (Anaphlebotomus) huberti Depaquit, Leger & Robert, 

2002

29 36

Phlebotomus (Anaphlebotomus) rodhaini Parrot, 1930 2, 3, 4, 7, 9, 12, 

13, 14*, 19, 21, 

22*, 23, 25, 27, 

36*, 39, 44, 48, 

50, 53, 55

3, 19, 44, 47, 

49, X

Phlebotomus (Anaphlebotomus) rousettus Davidson, 1981 48 3, 19

Phlebotomus (Anaphlebotomus) vincenti Randrianambinintsoa & 

Depaquit, 2013

29 36

Phlebotomus (Euphlebotomus) argentipes Annandale and Brunetti, 

1908

75 87, 98, 99, 101, 

103, 106, 107, 

113, 117, 120

3, 19

Phlebotomus (Euphlebotomus) autumnalis Artemiev, 1980 60 3, 19

Phlebotomus (Euphlebotomus) caudatus Artemiev, 1978 60 3, 19

Phlebotomus (Euphlebotomus) mesghalii Rashti & Nadim, 1970 60, 66 3, 19, 43

Phlebotomus (Euphlebotomus) nadimi Javadian, Jalali-Galousang & 

Seyedi-Rashti, 1997

66 3, 19, 43

Phlebotomus (Larroussius) aculeatus Lewis, Minter & Ashford, 1974 19, 25 3, 19, X

Phlebotomus (Larroussius) ariasi Tonnoir, 1921 1, 35, 54 95, 100, 108, 

112

3, 19, X

Phlebotomus (Larroussius) ashfordi Gebre-Michael and Lane, 1996 19 3, 19, 28, X

Phlebotomus (Larroussius) chadlii Rioux, Juminer and Gibily, 1966 1, 35, 54 3, 19

Phlebotomus (Larroussius) elgonensis Ngoka, Madel & Mutinga, 

1975

19, 25, 29 3, 19, 46, X

Phlebotomus (Larroussius) fantalensis Lewis, Minter & Ashford, 

1974

19 3, 19

Phlebotomus (Larroussius) gibiensis Lewis, Minter & Ashford, 1974 19 3, 19

Phlebotomus (Larroussius) guggisbergi Kirk & Lewis, 1952 25, 52, 55 3, 19

Phlebotomus (Larroussius) ilami Javadian, Jalali-Galousang & 

Seyedi-Rashti, 1997

66 43, 45

*New country species record. Notes: Africa includes several Western Indian Ocean islands (Comoros, Madagascar, Mauritius, Mayotte, Réunion, and 

Seychelles); X indicates observed.
January – June 2017 75

THE ARMY MEDICAL DEPARTMENT JOURNAL

Table 3 (continued; part 2 of 9). Checklist of Phlebotomine sand flies of Africa, Southwest and Central Asia, including distribution. 

Note: countries are represented by numerical codes which are defined in Table 4.

Species Africa* Southwest

Asia and

Central Asia*

Other

Countries

Selected

References

Phlebotomus (Larroussius) kandelakii kandelakii Shchurenkova, 

1929

60, 61, 62, 65, 

66, 73, 80

3, 19, 43

Phlebotomus (Larroussius) kandelakii burneyi Lewis, 1967 66, 75 3, 19

Phlebotomus (Larroussius) keshishiani Shchurenkova, 1936 60, 66, 75, 79 3, 19, 43

Phlebotomus (Larroussius) langeroni Nitzulescu, 1930 1, 35, 54 64, 66 3, 19

Phlebotomus (Larroussius) longicuspis Nitzulescu, 1930 1, 5, 28, 35, 54 112 3, 19, 25

Phlebotomus (Larroussius) longipes Parrot & Martin, 1939 19, 25, 50 3, 19, X

Phlebotomus (Larroussius) major Annandale, 1910 66, 75 107, 117 3, 19

Phlebotomus (Larroussius) mariae Rioux, Croset, Leger & BaillyChoumara, 1974

35 3, 19

Phlebotomus (Larroussius) neglectus Tonnoir, 1921 66, 80, 82 86, 88, 96, 100, 

110

3, 19, 43

Phlebotomus (Larroussius) notus Artemiev & Neronov, 1984 60 3, 19

Phlebotomus (Larroussius) orientalis Parrot, 1936 10, 15, 16, 19, 

25, 38, 41, 50, 

55

64, 77, 85 3, 19, 46, X

Phlebotomus (Larroussius) pedifer Lewis, Mutinga & Ashford, 1972 19, 25, 50 3, 19, X

Phlebotomus (Larroussius) perfiliewi perfiliewi Parrot, 1930 1,35, 54 62, 66, 80, 82 96, 97, 100, 

104, 109 

3, 19, 43

Phlebotomus (Larroussius) perfiliewi galilaeus Theodor, 1958 68, 80 94 3, 19

Phlebotomus (Larroussius) perfiliewi transcaucasicus Perfil'ev, 

1937

62, 65, 66. 67, 

80

3, 19

Phlebotomus (Larroussius) perniciosus Newstead, 1911 1, 28, 35 60, 65, 78, 80, 94, 95, 100, 

104, 108, 111, 

112

3, 19

Phlebotomus (Larroussius) smirnovi Perfil'ev, 1941 66, 70, 81, 84 92 3, 19

Phlebotomus (Larroussius) somaliensis Abonnenc, Adam & BaillyChoumara, 1959

19, 47 3, 19, X

Phlebotomus (Larroussius) syriacus Adler & Theodor, 1931 61, 62, 65, 68, 

69, 78, 80, 82

3, 19

Phlebotomus (Larroussius) tobbi Adler & Theodor, 1930 62, 65, 66, 67, 

68, 73, 78, 80 

86, 94, 96, 100 3, 19, 43

Phlebotomus (Larroussius) wenyoni Adler & Theodor, 1930 66, 67, 80, 81 3, 19, 43

Phlebotomus (Legeromyia) multihamatus Rahola, Depaquit, 

Makanga & Paupy, 2013

20 35

Phlebotomus (Madaphlebotomus) berentiensis (Leger & Rodhain, 

1978)

29 3, 19, 26, 33

Phlebotomus (Madaphlebotomus) fertei Depaquit, Leger & Robert, 

2002

29 21, 26, 36

Phlebotomus (Madaphlebotomus) fontenillei Depaquit, Leger & 

Robert, 2004

29 23, 26

Phlebotomus (Madaphlebotomus) vaomalalae Randrianambinintsoa, Leger & Depaquit, 2013

29 26, 37

Phlebotomus (Madaphlebotomus) vincenti Randrianambinintsoa & 

Depaquit, 2013

29 37

Phlebotomus (Paraphlebotomus) alexandri Sinton, 1928 1, 15, 16, 19, 

38, 50, 54

60, 62, 66, 67, 

68, 69, 70, 71, 

74, 75, 77, 80, 

81, 83, 85 

92, 94, 96, 98, 

105, 109, 112 

3, 19, 43, X

Phlebotomus (Paraphlebotomus) andrejevi Shakirzyanova, 1953 60, 66, 70, 84 92, 105 3, 19, 43

Phlebotomus (Paraphlebotomus) caucasicus Marzinowsky, 1917 60, 61, 62, 65, 

66, 70, 84 92 3, 19, 43

Phlebotomus (Paraphlebotomus) chabaudi Croset, Abonnenc & 

Rioux, 1970

1, 28, 35, 54 112 3, 19

*New country species record. Notes: Africa includes several Western Indian Ocean islands (Comoros, Madagascar, Mauritius, Mayotte, Réunion, and 

Seychelles); X indicates observed.
76 http://www.cs.amedd.army.mil/amedd_journal.aspx

NEW RECORDS, DISTRIBUTION, AND UPDATED CHECKLISTS OF OLD WORLD

PHLEBOTOMINE SAND FLIES, WITH EMPHASIS ON AFRICA, SOUTHWEST ASIA, AND CENTRAL ASIA

Table 3 (continued; part 3 of 9). Checklist of Phlebotomine sand flies of Africa, Southwest and Central Asia, including distribution. 

Note: countries are represented by numerical codes which are defined in Table 4.

Species Africa* Southwest

Asia and

Central Asia*

Other

Countries

Selected

References

Phlebotomus (Paraphlebotomus) gemetchi Gebre-Michael and 

Balkew, 2003

19 29

Phlebotomus (Paraphlebotomus) jacusieli Theodor, 1947 73*, 82, 66, 68, 

80

3, 19, 43, X

Phlebotomus (Paraphlebotomus) kazeruni Theodor & Mesghali, 

1964

1, 16, 27, 35 60, 66, 69, 77, 

80 

3, 19, 43, 47, X

Phlebotomus (Paraphlebotomus) marismortui Theodor, 1947 68 3, 19

Phlebotomus (Paraphlebotomus) mireillae Kellick-Kendrick, Tang, 

Johnson, Ngumbi & Robert, 1997

19, 25 46, X

Phlebotomus (Paraphlebotomus) mongolensis Sinton, 1928 60, 62, 66, 70, 

84

92 3, 19, 43

Phlebotomus (Paraphlebotomus) nuri Lewis, 1967 60, 66, 75 3, 19

Phlebotomus (Paraphlebotomus) saevus Parrot & Martin, 1939 7, 19, 25, 50 77, 85 3, 19

Phlebotomus (Paraphlebotomus) sergenti sergenti Parrot, 1917 1, 15, 16, 18, 19, 

25, 27, 28, 31, 35, 

37, 38, 47, 54

60, 61 64, 65, 

66, 67, 68, 69, 

70, 73, 75, 77, 

78, 80, 85 

94, 95, 96, 98, 

100, 104, 107, 

108, 109, 111, 

112

3, 19, 43, 47, X

Phlebotomus (Paraphlebotomus) sergenti similis Perfil'ev, 1963 63, 66, 82 118, 119 3, 19, 43

Phlebotomus (Phlebotomus) bergeroti Parrot, 1934 1, 5, 10, 15, 16, 

18, 19, 32, 35, 38, 

44, 47, 50

66, 68, 69, 75, 

77, 83, 85

3, 19, 43, 46, X

Phlebotomus (Phlebotomus) duboscqi Neveu-Lemaire, 1906 5, 7, 9, 10, 19, 21, 

22, 25, 31, 32, 33, 

38, 39, 44, 46, 

50, 53, 59

77, 85 3, 19, 40, 46, X

Phlebotomus (Phlebotomus) gigas Parrot & Schwetz, 1937 2, 7, 9, 12, 13, 

20, 23, 31

3, 19, 46, X

Phlebotomus (Phlebotomus) minteri Lewis, 1982 52 3, 19

Phlebotomus (Phlebotomus) papatasi (Scopoli, 1786) 1, 16, 18, 19, 25*, 

28, 35, 38, 47, 

50, 54

60, 61, 62, 65, 

66. 67. 68, 69, 

70, 71, 73, 74, 

75, 77, 78, 80, 

81, 84, 85

87, 89, 94, 96, 

96, 97, 98, 100, 

104, 107, 109, 

112, 113

3, 19, 42, 43, X

Phlebotomus (Phlebotomus) salehi Mesghali, 1965 66 98 3, 19, 43, X

Phlebotomus (Synphlebotomus) ansarii Lewis, 1957 66 3, 19, 43

Phlebotomus (Synphlebotomus) celiae Minter, 1962 19, 25 46, X

Phlebotomus (Synphlebotomus) eleanorae Sinton, 1931 66 98 3, 19, 43

Phlebotomus (Synphlebotomus) grovei Downes, 1971 37 3, 19

Phlebotomus (Synphlebotomus) katangensis Bequaert & Walravens, 1930

12, 58 3, 19, 46, X

Phlebotomus (Synphlebotomus) martini Parrot, 1936 19, 25, 47, 50, 55 3, 19, X

Phlebotomus (Synphlebotomus) rossi de Meillon & Lavoipierre, 

1944

37, 48, 58 3, 19

Phlebotomus (Synphlebotomus) saltiae Leger, Haddad and 

Chaker, 1997

73 32

Phlebotomus (Synphlebotomus) taylori Davidson, 1982 58 3, 19

Phlebotomus (Synphlebotomus) vansomerenae Heisch, Guggisberg & Teesdale, 1956

19, 25 3, 19

Phlebotomus (Transphlebotomus) canaaniticus Adler & Theodor, 

1931

68, 69, 78 3, 19

Phlebotomus (Transphlebotomus) mascittii Grassi, 1908 1 6 94, 95, 96, 100, 

102, 112, 115

3, 19

Sergentomyia (Capensomyia) caffrarica (de Meillon & Lavoipierre, 

1944)

48 3, 19

Sergentomyia (Capensomyia) capensis (de Meillon, 1955) 48 3, 19

*New country species record. Notes: Africa includes several Western Indian Ocean islands (Comoros, Madagascar, Mauritius, Mayotte, Réunion, and 

Seychelles); X indicates observed.
January – June 2017 77

THE ARMY MEDICAL DEPARTMENT JOURNAL

Table 3 (continued; part 4 of 9). Checklist of Phlebotomine sand flies of Africa, Southwest and Central Asia, including distribution. 

Note: countries are represented by numerical codes which are defined in Table 4.

Species Africa* Southwest

Asia and

Central Asia*

Other

Countries

Selected

References

Sergentomyia (Capensomyia) drakensbergi Davidson, 1979 48 3, 19

Sergentomyia (Capensomyia) haeselbarthi (Abonnenc, 1967) 48 3, 19

Sergentomyia (Capensomyia) kalaharia Davidson, 1979 37, 48 3, 19

Sergentomyia (Capensomyia) luteola Davidson, 1983 48 3, 19

Sergentomyia (Capensomyia) meeseri (de Meillon & Hard, 1953) 48, 55, 58 3, 19

Sergentomyia (Capensomyia) nama Davidson, 1953 37, 48 3, 19, X

Sergentomyia (Capensomyia) namibensis (de Meillon & Hardy, 1953) 37, 48 3, 19, X

Sergentomyia (Capensomyia) xera Davidson, 1979 37, 48 3, 19, X

Sergentomyia (Demeillonius) transvaalensis Sinton, 1933 48 3, 19

Sergentomyia (Grassomyia) dreyfussi dreyfussi (Parrot, 1933) 1, 15, 18, 19, 

25, 35, 47, 54, 

66 

66, 69, 77, 85 3, 19, 43

Sergentomyia (Grassomyia) dreyfussi turkestanica Theodor & 

Mesghali, 1964

60, 68, 81 3, 19

Sergentomyia (Grassomyia) ghesquierei (Parrot, 1929) 3, 5, 12, 13, 14, 

19, 21, 22*, 23, 

44, 47*

3, 19, X

Sergentomyia (Grassomyia) indica (Theodor, 1931) 66, 75 90, 92, 93, 98, 

99, 101, 103, 

107, 116, 117

3, 19

Sergentomyia (Grassomyia) inermis (Theodor, 1938) 2, 3, 4, 5, 9, 10, 

19, 21, 22, 23, 

25, 30, 39, 44, 

48, 50

3, 19, 49

Sergentomyia (Grassomyia) madagascariensis (Abonnenc, 1969) 29 3, 19

Sergentomyia (Grassomyia) squamipleuris (Newstead, 1912) 2, 3, 5, 9, 10, 

12, 13, 14, 15, 

16, 19, 21, 22, 

23, 25, 29, 30, 

31, 36, 39, 44, 

46, 48, 50, 52, 

53, 55, 57 

66, 67, 68, 77 3, 19, 27, 43, X

Sergentomyia (Neophlebotomus) angolensis (Abonnenc, 1968) 2, 13 3, 19

Sergentomyia (Neophlebotomus) collarti (Adler, Theodor & Parrot, 

1929)

2, 7, 9, 12, 13, 

14, 21, 22, 23, 

39, 42, 46, 50, 

52, 55

3, 19, X

Sergentomyia (Neophlebotomus) corneti Pastre, 1975 44 3, 19, X

Sergentomyia (Neophlebotomus) decipiens (Theodor, 1931) 7, 9, 12, 13, 14, 

23, 25, 39, 44, 

50, 52*, 55

3, 19, X

Sergentomyia (Neophlebotomus) dolichopa (Abonnenc & Courtois, 

1970)

15 66, 85 3, 19, X

Sergentomyia (Neophlebotomus) dureni Parrot, 1934 2, 5, 9, 10, 12, 

13, 14, 19*, 21, 

22*, 23, 25, 27, 

39, 44, 50, 52

3, 19, X

Sergentomyia (Neophlebotomus) dyemkoumai (Abonnenc, 1964) 13, 14 3, 19, 46

Sergentomyia (Neophlebotomus) grejbinei (Vattier-Bernard, 1971) 12, 13 3, 19

Sergentomyia (Neophlebotomus) grilloti (Vattier-Bernard & Bimangou, 1975)

13 3, 19

Sergentomyia (Neophlebotomus) harveyi (Heisch, Guggisberg & 

Teesdale), 1956

25 3, 19

Sergentomyia (Neophlebotomus) hodgsoni (Sinton, 1933) 60, 66, 75 98, 117 3, 19

*New country species record. Notes: Africa includes several Western Indian Ocean islands (Comoros, Madagascar, Mauritius, Mayotte, Réunion, and 

Seychelles); X indicates observed.
78 http://www.cs.amedd.army.mil/amedd_journal.aspx

NEW RECORDS, DISTRIBUTION, AND UPDATED CHECKLISTS OF OLD WORLD

PHLEBOTOMINE SAND FLIES, WITH EMPHASIS ON AFRICA, SOUTHWEST ASIA, AND CENTRAL ASIA

Table 3 (continued; part 5 of 9). Checklist of Phlebotomine sand flies of Africa, Southwest and Central Asia, including distribution. 

Note: countries are represented by numerical codes which are defined in Table 4.

Species Africa* Southwest

Asia and

Central Asia*

Other

Countries

Selected

References

Sergentomyia (Neophlebotomus) ingrami (Newstead, 1914) 2, 3, 5, 7, 9, 

12, 13, 14, 19*, 

21, 22, 23, 25, 

27, 39, 44, 50, 

53, 55

19, 47, X

Sergentomyia (Neophlebotomus) kirki (Parrot, 1948) 25, 30, 50 3, 19

Sergentomyia (Neophlebotomus) kitonyii (Minter, 1963) 19, 25 3, 19, X

Sergentomyia (Neophlebotomus) machadoi (Abonnenc, 1968) 2 3, 19, X

Sergentomyia (Neophlebotomus) macintoshi (Abonnenc & Pastre, 

1972)

48 3, 19

Sergentomyia (Neophlebotomus) notata (Parrot, 1938) 19 3, 19

Sergentomyia (Neophlebotomus) pawlowskyi Perfil'ev, 1933 60, 61, 66, 67, 

80, 81, 84

3, 19

Sergentomyia (Neophlebotomus) serrata (Parrot & Malbrant, 1945) 12, 13, 19, 25, 

50, 55

3, 19

Sergentomyia (Neophlebotomus) sonyae Lewis, 1982 74, 77 3, 19

Sergentomyia (Neophlebotomus) teesdalei Minter, 1963 25 3, 19, X

Sergentomyia (Neophlebotomus) trouilleti Vattier-Bernard, 1976 13 3, 19

Sergentomyia (Parrotomyia) africana africana (Newstead, 1912) 3, 5, 7, 9, 12, 

13, 15, 14, 19, 

21, 22,23, 25, 

27, 31, 32, 38, 

39, 44, 46, 48, 

50, 53, 55, 57, 

85

66 3, 27, 34, 43, 

46, 47, X

Sergentomyia (Parrotomyia) africana asiatica (Theodor, 1933) 35 68, 69, 75, 77 98 3, 19

Sergentomyia (Parrotomyia) babu babu (Annandale, 1910) 33 60, 75 87, 98 3, 19

Sergentomyia (Parrotomyia) baghdadis Adler & Theodor, 1929 60, 66, 67, 75 98 3, 19, 43, X

Sergentomyia (Parrotomyia) bailyi (Sinton, 1931) 66, 75 90, 92, 98, 101, 

117, 120

3, 19

Sergentomyia (Parrotomyia) crosarai (Parrot & Wanson, 1946) 2, 12, 13, 27 3, 19, 47, X

Sergentomyia (Parrotomyia) eremitis (Parrot & Bouquet de Joliniere, 

1945)

1, 27, 50, 53 3, 19, 47, X

Sergentomyia (Parrotomyia) freetownensis freetownensis (Sinton, 

1930)

2, 3, 9, 14, 18, 

12*, 13, 19, 25, 

23, 32, 44, 46, 

48, 50, 58, 77

3, 19, X

Sergentomyia (Parrotomyia) freetownensis furanus (Lewis & Kirk, 

1958)

50 3, 19

Sergentomyia (Parrotomyia) grekovi Khodukin, 1929 60, 66, 70, 75, 

79, 81, 82, 84

98 3, 19, 43

Sergentomyia (Parrotomyia) magna (Sinton, 1932) 2, 5, 7, 9, 12, 

14, 21, 19, 22*, 

23, 31, 32, 39, 

53, 44, 48, 50, 

58

77 3, 19, X

Sergentomyia (Parrotomyia) palestinensis (Adler & Theodor, 1927) 1, 15, 16, 18, 

19, 35, 50

62, 66, 67, 68, 

69, 75, 77

98 3, 19, 43, X

Sergentomyia (Parrotomyia) rhodesiensis (de Meillon & Hardy, 

1953)

58 3, 19

Sergentomyia (Parrotomyia) shorttii (Adler & Theodor, 1927) 75 87, 98, 106 3, 19

Sergentomyia (Parrotomyia) sogdiana (Parrot, 1928) 66, 79, 84 3, 19, 43

Sergentomyia (Parrotomyia) sumbarica Perfil'ev, 1933 60, 66, 67, 81, 

84

92 3, 19, 43, X

Sergentomyia (Parrotomyia) yvonnae (Parrot & Schwetz, 1937) 12 3, 19, X

*New country species record. Notes: Africa includes several Western Indian Ocean islands (Comoros, Madagascar, Mauritius, Mayotte, Réunion, and 

Seychelles); X indicates observed.
January – June 2017 79

THE ARMY MEDICAL DEPARTMENT JOURNAL

Table 3 (continued; part 6 of 9). Checklist of Phlebotomine sand flies of Africa, Southwest and Central Asia, including distribution. 

Note: countries are represented by numerical codes which are defined in Table 4.

Species Africa* Southwest

Asia and

Central Asia*

Other

Countries

Selected

References

Sergentomyia (Parrotomyia) zielkei Seccombe, Ready and Huddleston, 1993

48 3, 19

Sergentomyia (Parvidens) arida Davidson, 1982 37, 58 3, 19, X

Sergentomyia (Parvidens) heischi Kirk & Lewis, 1950 19, 25, 36, 37, 

50

3, 19, X

Sergentomyia (Parvidens) iranica (Lewis & Mesghali, 1961) 66 3, 19, 43

Sergentomyia (Parvidens) lesleyae (Lewis & Kirk, 1946) 19, 32, 50 3, 19

Sergentomyia (Rondanomyia) goodmani goodmani Depaquit & 

Robert, 2005

29 22

Sergentomyia (Rondanomyia) goodmani comorensis Depaquit, 

Randrianamnini & Leger, 2012

11 39

Sergentomyia (Sergentomyia) antennata (Newstead, 1912) 1, 3, 5, 7, 9, 

10, 19, 12, 14, 

15, 16, 19, 20, 

21, 22, 23, 25, 

27, 30, 31, 32, 

35, 39, 44, 46, 

47, 48, 50, 52*, 

53, 54

66, 67, 68, 69, 

71, 77, 83, 55

3, 19, 43, 46, 

47, X

Sergentomyia (Sergentomyia) arpaklensis Perfil'ev, 1933 81, 84 3, 19

Sergentomyia (Sergentomyia) ashfordi Davidson, 1987 19 3, 19

Sergentomyia (Sergentomyia) bedfordi (Newstead, 1914) 2, 3, 4**, 7, 9, 

10, 12, 14, 19, 

21, 22, 23,25, 

27,31, 36, 37, 

39, 44, 48, 50, 

52, 53, 55, 58

3, 19, 47, 49, X

Sergentomyia (Sergentomyia) bergerardi Trouillet & Vattier-Bernard, 

1978

13 3, 19

Sergentomyia (Sergentomyia) bimangoui Davidson, 1990 13 3, 19

Sergentomyia (Sergentomyia) blossi (Kirk & Lewis, 1952) 25 3, 19, X

Sergentomyia (Sergentomyia) buxtoni (Theodor, 1933) 3, 5, 10, 14*, 

21, 22, 31, 38, 

39, 44

3, 19, X

Sergentomyia (Sergentomyia) caliginosa Davidson, 1987 30, 36, 48, 57, 

58

3, 19

Sergentomyia (Sergentomyia) cincta (Parrot & Martin, 1944) 9, 15, 16, 19, 

22, 25, 50, 55

3, 19, X

Sergentomyia (Sergentomyia) congolensis (Bequaert &Walravens, 

1930)

4, 12 3, 49, X

Sergentomyia (Sergentomyia) davidsoni Seccombe, Ready and 

Huddleston, 1993

18, 19, 50 3, 19, X

Sergentomyia (Sergentomyia) dentata dentata (Sinton, 1933) 60, 62, 65, 66, 

67, 75, 78, 80, 

83, 85

3, 19, 43

Sergentomyia (Sergentomyia) dentata agdamica Artemiev, 1982 61, 62, 65, 66, 

67

3, 19

Sergentomyia (Sergentomyia) dentata bruchoni Parrot, 1935 80 96 3, 19

Sergentomyia (Sergentomyia) dubia (Parrot, Mornet & Cadenat, 

1934)

5, 7, 13, 14, 21, 

22, 23, 31, 32, 

39, 44

3, 19, X

Sergentomyia (Sergentomyia) fallax fallax Parrot, 1921 1, 5, 15, 16, 19, 

35, 38, 54

60, 64, 77, 85 3, 19, X

Sergentomyia (Sergentomyia) fallax afghanica Artemiev, 1974 60 3, 19

Sergentomyia (Sergentomyia) fallax cypriotica (Adler, 1946) 67, 68, 73, 80 94 3, 19

Sergentomyia (Sergentomyia) formica Davidson, 1987 37, 48, 58 3, 19

Sergentomyia (Sergentomyia) gracilis (Kirk & Lewis, 1952) 25 3, 19

*New country species record; **Sergentomyia (Sergentomyia) bedfordi “Maun form”. Notes: Africa includes several Western Indian Ocean islands (Comoros, 

Madagascar, Mauritius, Mayotte, Réunion, and Seychelles); X indicates observed.
80 http://www.cs.amedd.army.mil/amedd_journal.aspx

NEW RECORDS, DISTRIBUTION, AND UPDATED CHECKLISTS OF OLD WORLD

PHLEBOTOMINE SAND FLIES, WITH EMPHASIS ON AFRICA, SOUTHWEST ASIA, AND CENTRAL ASIA

Table 3 (continued; part 7 of 9). Checklist of Phlebotomine sand flies of Africa, Southwest and Central Asia, including distribution. 

Note: countries are represented by numerical codes which are defined in Table 4.

Species Africa* Southwest

Asia and

Central Asia*

Other

Countries

Selected

References

Sergentomyia (Sergentomyia) impudica Abonnenc, 1968 2, 13, 27, 55 3, 19, 44, X

Sergentomyia (Sergentomyia) logonensis (Rageau, 1951) 7, 9 3, 19, X

Sergentomyia (Sergentomyia) magnidentata Davidson, 1987 25, 47* 3, 19, X

Sergentomyia (Sergentomyia) mervynae Pringle, 1953 54 60, 66, 75 3, 19, 43

Sergentomyia (Sergentomyia) minuta minuta Rondani, 1843 25* 80, 82 95, 96, 100, 

104, 108, 109, 

112

3, 19, X

Sergentomyia (Sergentomyia) minuta parroti (Adler & Theodor, 

1927)

1, 28, 35, 54 78 94, 96, 112 3, 19

Sergentomyia (Sergentomyia) murgabiensis murgabiensis Perfil'ev, 

1939

60, 66, 70, 81 3, 19

Sergentomyia (Sergentomyia) murgabiensis pashtunica Artemiev, 

1974

60, 66, 75 105 3, 19

Sergentomyia (Sergentomyia) murgabiensis sintoni Pringle, 1953 16 60, 66, 67, 71, 

80, 81

3, 19

Sergentomyia (Sergentomyia) pastoriana (Parrot, Mornet & Cadenat, 1945)

14, 19, 22, 23, 

39, 46, 50, 55

3, 19, X

Sergentomyia (Sergentomyia) punjabensis (Sinton, 1933) 75 113, 117 3, 19

Sergentomyia (Sergentomyia) renauxi (Parrot & Schwetz, 1937) 12, 13 3, 19, X

Sergentomyia (Sergentomyia) richardi (Parrot & Wanson, 1946) 7, 12, 27, 53, 55 3, 19, X

Sergentomyia (Sergentomyia) rima Davidson, 1987 30, 36, 37, 48, 

57, 58

3, 19, X

Sergentomyia (Sergentomyia) ruttledgei (Lewis & Kirk, 1946) 50, 55 3, 19

Sergentomyia (Sergentomyia) salisburiensis (Abonnenc, 1967) 4, 37, 48, 49, 

57, 58

3, 19, 49, X

Sergentomyia (Sergentomyia) schoutedeni schoutedeni (Adler, 

Theodor & Parrot, 1929)

3, 7, 12, 14, 23, 

25, 26, 39, 50, 

53, 55

3, 19, X

Sergentomyia (Sergentomyia) schoutedeni nocens (Parrot, 1951) 25, 50 3, 19

Sergentomyia (Sergentomyia) schwetzi (Adler, Theodor & Parrot, 

1929)

1, 2, 3, 5, 7, 9, 

10, 12, 13, 14, 

15, 16, 19, 20, 

21, 22, 23, 25, 

27, 31, 32, 35, 

35, 37, 39, 44, 

46, 47, 48, 50, 

52*, 53, 55

77, 85 24, 25, 27, X

Sergentomyia (Sergentomyia) serridentata Davidson, 1990 7, 55 3, 19, X

Sergentomyia (Sergentomyia) silva Trouillet, 1985 13 3, 19

Sergentomyia (Sergentomyia) taizi Lewis, 1974 15, 16 77, 85 3, 19, X

Sergentomyia (Sergentomyia) teteica Artemiev, 1985 36 3, 19

Sergentomyia (Sergentomyia) theodori (Parrot, 1919) 16 60, 66, 67, 68, 

69, 73, 75, 77, 

78, 80

94, 96, 98, 101 3, 19, 43

Sergentomyia (Sergentomyia) waqqasi Artemiev, 1982 67 3, 19

Sergentomyia (Sergentomyia) yusafi (Sinton, 1930) 25, 39, 52, 55, 

57

85 3, 19, 46, X

Sergentomyia (Sergentomyia) zumpti (Abonnenc, 1967) 37, 48 3, 19

Sergentomyia (Sintonius) adami (Abonnenc, 1960) 9, 31 3, 19, X

Sergentomyia (Sintonius) adleri (Theodor, 1933) 3, 5, 7, 9, 10, 

14, 15, 16, 19, 

21, 22, 25, 38, 

39, 44, 50, 53, 

55

66, 69, 77, 83 3, 19, 27, 46, X

*New country species record. Notes: Africa includes several Western Indian Ocean islands (Comoros, Madagascar, Mauritius, Mayotte, Réunion, and 

Seychelles); X indicates observed.
January – June 2017 81

THE ARMY MEDICAL DEPARTMENT JOURNAL

Table 3 (continued; part 8 of 9). Checklist of Phlebotomine sand flies of Africa, Southwest and Central Asia, including distribution. 

Note: countries are represented by numerical codes which are defined in Table 4.

Species Africa* Southwest

Asia and

Central Asia*

Other

Countries

Selected

References

Sergentomyia (Sintonius) affinis affinis (Theodor, 1933) 9, 19, 23, 25, 50 3, 19, 46, X

Sergentomyia (Sintonius) affinis vorax (Parrot, 1948) 2, 3, 5, 9, 10, 

19, 22, 23, 25, 

31, 39, 44, 50

3, 19, X

Sergentomyia (Sintonius) balmicola Abonnenc, Adam & BaillyChoumara, 1959

7 3, 19

Sergentomyia (Sintonius) calcarata (Parrot, 1948) 10, 15, 18, 19, 

50

77 3, 19, X

Sergentomyia (Sintonius) choumarai (Abonnenc, 1960) 47 3, 19

Sergentomyia (Sintonius) christophersi (Sinton, 1927) 1, 5, 9, 10, 15, 

16, 18, 19, 21, 

23, 25, 29, 35, 

38, 39, 44, 50, 

54, 55

60, 66, 69, 74, 

75, 77, 85

98 3, 19, 27, 43, 

46, X

Sergentomyia (Sintonius) clastrieri (Abonnenc, 1964) 15, 23 3, 19

Sergentomyia (Sintonius) clydei (Sinton, 1928) 1, 5, 7, 9, 10, 

15, 16, 18, 19, 

21, 22, 25, 31, 

35, 38, 39, 44, 

45, 47, 50, 53, 

54, 55

60, 66, 67, 68, 

70, 71, 75, 77, 

79, 81, 83, 85

3, 19, 44, X

Sergentomyia (Sintonius) diapagai (Abonnenc, 1962) 5, 31 3, 19

Sergentomyia (Sintonius) edentulus Pastre, 1982 44 3, 19, X

Sergentomyia (Sintonius) graingeri (Heisch, Guggisberg & Teesdale, 

1956)

25 3, 19

Sergentomyia (Sintonius) herollandi (Abonnenc, 1960) 31, 44 3, 19, X

Sergentomyia (Sintonius) hospitii (Sinton, 1924) 75 98 3, 19

Sergentomyia (Sintonius) lewisiana Pastre, 1982 44 3, 19, X

Sergentomyia (Sintonius) mbandakai Abonnenc, 1970 12 3, 19

Sergentomyia (Sintonius) meilloni Sinton, 1932 25, 36, 37, 48, 

51, 58

3, 19, X

Sergentomyia (Sintonius) rogeri Pastre, 1982 44 3, 19, X

Sergentomyia (Sintonius) sattii Qutubuddin, 1962 50 3, 19

Sergentomyia (Sintonius) sidioliensis Pastre, 1982 44 3, 19, X

Sergentomyia (Sintonius) suberecta (Sinton, 1932) 25, 50 3, 19, X

Sergentomyia (Sintonius) tauffliebi (Abonnenc & Cornet, 1971) 13, 14, 44 3, 19, X

Sergentomyia (Sintonius) thomsoni (Theodor, 1938) 10, 19, 25, 30 3, 19

Sergentomyia (Sintonius) tiberiadis tiberiadis (Adler, Theodor & 

Lourie, 1930)

15, 16, 18, 19, 

50

60, 66, 68, 74, 

77, 83, 85

3, 19, 43, X

Sergentomyia (Sintonius) tiberiadis pakistanica Artemiev & 

Saf'janova, 1974

60, 75, 81 3, 19

Sergentomyia (Sintonius) wansoni (Parrot, 1938) 2, 5, 9, 12, 13, 

19, 31, 55

66 3, 19, X

Sergentomyia (Spelaeomyia) bailyi (Sinton, 1931) 7 75 92, 98, 117, 

120

3, 19

Sergentomyia (Spelaeomyia) darlingi (Lewis & Kirk, 1954) 5, 9, 31,50 3, 19, X

Sergentomyia (Spelaeomyia) emilii (Vattier-Bernard, 1966) 5, 9, 13 3, 19, X

Sergentomyia (Spelaeomyia) hunti (Lewis & Kirk, 1946) 9, 50 3, 19

Sergentomyia (Spelaeomyia) mirabilis (Parrot & Wanson, 1939) 2, 9, 12, 13, 

52, 55

3, 19, X

Sergentomyia (Spelaeomyia) moucheti (Vattier-Bernard & Abonnenc, 1967)

7, 9, 13 3, 19

*New country species record. Notes: Africa includes several Western Indian Ocean islands (Comoros, Madagascar, Mauritius, Mayotte, Réunion, and 

Seychelles); X indicates observed.
82 http://www.cs.amedd.army.mil/amedd_journal.aspx

NEW RECORDS, DISTRIBUTION, AND UPDATED CHECKLISTS OF OLD WORLD

PHLEBOTOMINE SAND FLIES, WITH EMPHASIS ON AFRICA, SOUTHWEST ASIA, AND CENTRAL ASIA

Table 3 (continued; part 9 of 9). Checklist of Phlebotomine sand flies of Africa, Southwest and Central Asia, including distribution. 

Note: countries are represented by numerical codes which are defined in Table 4.

Species Africa* Southwest

Asia and

Central Asia*

Other

Countries

Selected

References

Sergentomyia (Trouilletomyia) boironis Ranadrianambinintspa & 

Depaquit, 2014 29 38

Sergentomyia (Trouilletomyia) huberti Depaquit, Leger & Robert, 

2002

29 24

Sergentomyia (Vattieromyia) anka Depaquit, Leger & Robert, 2008 29 24

Sergentomyia (Vattieromyia) namo Depaquit, Leger & Robert, 2008 29 24

Sergentomyia (Vattieromyia) pessoni Depaquit, Randrianamnini & 

Leger, 2012

11 39

Sergentomyia (Vattieromyia) schlerpsiphon Depaquit, Leger & 

Robert, 2008

29 24

Sergentomyia (Ungrouped)** berentiensis Leger & Rodhain, 1978 29 3, 19

Sergentomyia (Ungrouped) bernardae Trouillet, 1982 13 3, 19

Sergentomyia (Ungrouped) cunicula Davidson, 1979 37 3, 19

Sergentomyia (Ungrouped) dissimillima (Abonnenc, 1972) 5, 13, 14, 23 3, 19

Sergentomyia (Ungrouped) garnhami (Heisch, Guggisberg & Teesdale, 1956)

25, 52 3, 19

Sergentomyia (Ungrouped) hamoni (Abonnenc, 1958) 2, 5, 7, 9, 12, 

13, 14, 31, 44, 

50

3, 19, X

Sergentomyia (Ungrouped) hirta (Parrot & Bouquet de Joliniere, 

1945)

1 3, 19

Sergentomyia (Ungrouped) horridula Vattier-Bernard & Trouillet, 

1982

13 3, 19

Sergentomyia (Ungrouped) leponti Vattier-Bernard, 1973 13 3, 19

Sergentomyia (Ungrouped) lumsdeni Kirk & Lewis, 1950 13, 55 3, 19

Sergentomyia (Ungrouped) majungaensis Depaquit, Leger and 

Robert, 2007

29 48

Sergentomyia (Ungrouped) metzi Davidson, 1979 37, 48 3, 19, X

Sergentomyia (Ungrouped) montana (Sinton, 1924) 75 98, 107 3, 19

Sergentomyia (Ungrouped) moreli (Abonnenc & Hamon, 1958) 13, 14, 27, 55 3, 43, 47, X

Sergentomyia (Ungrouped) multidens (Heisch, Guggisberg & Teesdale, 1956)

15, 19, 25 85 3, 19

Sergentomyia (Ungrouped) ovazzai (Pastre, 1973) 23 3, 19

Sergentomyia (Ungrouped) roberti Vattier-Bernard & Trouillet, 1081 13 3, 19

Sergentomyia (Ungrouped) rosannae (Heisch, Guggisberg & Teesdale, 1956)

25 3, 19

Sergentomyia (Ungrouped) simillima (Newstead, 1914) 3, 5, 7, 9, 12, 

13, 14, 19, 22, 

23, 24, 27, 37, 

39, 46, 50, 53, 

55

3, 43, 47, X

Sergentomyia (Ungrouped) villosa Davidson, 1979 37, 48, 58 3, 19, X

Sergentomyia (Ungrouped) vulpes Davidson, 1979 37, 48 3, 19, X

Sergentomyia (Ungrouped) welwitschii Davidson, 1979 37, 48 3, 19, X

Sergentomyia (Ungrouped) wurtzi (Parrot, 1938) 19 3, 19

Sergentomyia (Ungrouped) wynnae (Watson, 1951) 25, 55 3, 19

*New country species record. Notes: Africa includes several Western Indian Ocean islands (Comoros, Madagascar, Mauritius, Mayotte, Réunion, and 

Seychelles); X indicates observed.

** Ungrouped indicates subgenus uncertain.
January – June 2017 83

THE ARMY MEDICAL DEPARTMENT JOURNAL

Table 4. List of countries and their numerical codes as shown in Figure 1 and Table 3.

Code

No.

Country* Code 

No.

Country* Code

No.

Country* Code 

No.

Country*

1 Algeria 31 Mali 62 Azerbaijan 92 China (mainland)

2 Angola 32 Mauritania 63 Bahrain 93 China (Hong Kong)

3 Benin 33 Mauritius 64 Egypt 94 Cyprus

4 Botswana 34 Mayotte (France) 65 Georgia 95 France

5 Burkina Faso 35 Morocco

(excluding Western Sahara) 66 Iran 96 Greece

6 Burundi 36 Mozambique 67 Iraq 97 Hungary

7 Cameroon 37 Namibia 68 Israel 98 India

8 Cape Verde 38 Niger 69 Jordan 99 Indonesia

9 Central African Republic 39 Nigeria 70 Kazakhstan 100 Italy

10 Chad 40 Réunion (France) 71 Kuwait 101 Laos

11 Comoros 41 Rwanda 72 Kyrgyzstan 102 Macedonia

12 Congo, Democratic

Republic of 42 Saint Helena, Ascension and Tristan da Cunha (UK) 73 Lebanon 103 Malaysia

13 Congo, Republic of 43 São Tomé and Príncipe 74 Oman 104 Malta

14 Côte d'Ivoire 44 Senegal 75 Pakistan 105 Mongolia

15 Djibouti 45 Seychelles 76 Qatar 106 Myanmar

16 Egypt 46 Sierra Leone 77 Saudi Arabia 107 Nepal

17 Equatorial Guinea 47 Somalia 78 Syria 108 Portugal

18 Eritrea 48 South Africa 79 Tajikistan 109 Romania

19 Ethiopia 49 South Sudan 80 Turkey 110 Russia

20 Gabon 50 Sudan 81 Turkmenistan 111 Serbia & Montenegro

(former Yugoslavia)

21 Gambia 51 Swaziland 82 Ukraine 112 Spain

22 Ghana 52 Tanzania 83 United Arab Emirates 113 Sri Lanka

23 Guinea 53 Togo 84 Uzbekistan

(including Crimea) 114 Sri Lanka (Ubduam)

24 Guinea-Bissau 54 Tunisia 85 Yemen 115 Switzerland

25 Kenya 55 Uganda 86 Albania 116 Taiwan

26 Lesotho 56 Western Sahara 87 Bangladesh 117 Thailand

27 Liberia 57 Zambia 88 Bosnia & Herzegovina 118 Ukraine

28 Libya 58 Zimbabwe 89 Bulgaria 119 Ukraine (Crimea)

29 Madagascar 60 Afghanistan 90 Cambodia 120 Vietnam

30 Malawi 61 Armenia 91 Cameroon

*Africa (code nos. 1-58, including West Indian Ocean Islands); Southwest Asia and Central Asia (code nos. 60-85, including Middle East/CENTCOM; part of 

EUCOM, former Russian Republics); other countries (Code nos. 86-121)

References

1. Sand Fly-Borne Diseases [internet]. CVBD Companion 

Vector-borne Diseases website. Bayer HealthCare AG, 

Leverkusen, Germany. Available at: http://www.cvbd.

org/en/sand-fly-borne-diseases/. Accessed October 11, 

2015.

2. Lane R. Sandflies (Phlebotominae). In: Lane RP, Crosskey RW, eds. Medical Insects and Arachnids. London, 

U.K: Chapman & Hall; 1993:78-119.

3. Seccombe AK, Ready PD, Huddleston LM, eds. A 

Catalogue of Old World Phlebotomine Sandflies (Diptera: Psychodidae, Phlebotominae). London, UK: Intercept; 1993. Issue 8, Occasional Papers on Systematic 

Entomology.

4. Young DG, Duncan MA. Guide to the identification 

and geographic distribution of Lutzomyia sand flies in 

Mexico, the West Indies, Central and South America 

(Diptera: Psychodidae). Gainsville, FL: Associated 

Publishers; 1994. Available at: http://www.dtic.mil/dtic/

tr/fulltext/u2/a285737.pdf. Accessed March 15, 2017.

5. Depaquit J, Grandadam M, Fouque F, Andry PE, 

Peyrefitte C. Arthropod-borne viruses transmitted by 

phlebotomine sandflies in Europe: a review. Eurosurveillance. 2010;15(10):1-8.

6. Tesh RB. The genus Phlebovirus and its vectors. Annu

Rev Entomol. 1998;33:169-181.

7. Ready PD. Biology of Phlebotomine sand flies as vectors 

of disease agents. Annu Rev Entomol. 2013;58:227-250.
84 http://www.cs.amedd.army.mil/amedd_journal.aspx

NEW RECORDS, DISTRIBUTION, AND UPDATED CHECKLISTS OF OLD WORLD

PHLEBOTOMINE SAND FLIES, WITH EMPHASIS ON AFRICA, SOUTHWEST ASIA, AND CENTRAL ASIA

8. Maroli M, Feliciangeli MD, Bichaud L, Charrel RN, 

Gradoni L. Phlebotomine sandflies and the spreading of 

leishmaniases and other diseases of public health concern. Med Vet Entomol. 2013;27:123-147.

9. Bates PA, Depaquit J, Galati EAB, et al. Recent advances in phlebotomine sand fly research related to leishmaniasis control. Parasit Vectors. 2015;8:131. doi: 10.1186/

s13071-015-0712-x.

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Authors

Dr Rueda is an Adjunct Scientist of the Smithsonian Institution and formerly Research Entomologist and Chief 

of the Walter Reed Biosystematics Unit, Entomology 

Branch, Walter Reed Army Institute of Research, located at the Smithsonian Institution, Museum Support 

Center, Suitland, Maryland.

Mr J. Pecor is a Museum Specialist of the Walter Reed 

Biosystematics Unit, Entomology Branch, Walter Reed 

Army Institute of Research, located at the Smithsonian 

Institution, Museum Support Center, Suitland, Maryland.

Mr Wolkoff is a graduate student at the University of 

Texas at Tyler, Tyler, Texas and formerly Research Assistant of the Walter Reed Biosystematics Unit, Entomology Branch, Walter Reed Army Institute of Research, 

located at the Smithsonian Institution, Museum Support 

Center, Suitland, Maryland.

Mr D. Pecor is a VectorMap Data Management Technician of the Walter Reed Biosystematics Unit, Entomology Branch, Walter Reed Army Institute of Research, 

located at the Smithsonian Institution, Museum Support 

Center, Suitland, Maryland.

Ms Benyamin is an Associate Scientist of BioReliance, 

Rockville, Maryland and formerly O-RISE Technical 

Research Assistant of the Walter Reed Biosystematics 

Unit, Entomology Branch, Walter Reed Army Institute 

of Research, located at the Smithsonian Institution, Museum Support Center, Suitland, Maryland.

Dr Boussès is an Entomologist at the Institut de Recherche pour le Développement (IRD), Maladies Infectieuses 

et Vecteurs: Ecologie, Génétique, Evolution et Contrôle 

(MIVEGEC), Montpellier, France.

Dr Debboun is the Director of the Mosquito & Vector 

Control Division, Harris County Public Health, Houston, 

Texas.
86 http://www.cs.amedd.army.mil/amedd_journal.aspx

Vector-borne and zoonotic diseases pose a threat to active military personnel both overseas and in the United 

States. Deaths among service members from vectorborne diseases have dropped in recent decades, but 

morbidity from vector-borne diseases and injuries from 

stinging arthropods remain a real threat.1-4 The US Air 

Force (USAF) maintains arthropod identification and 

pathogen surveillance capability at the USAF School 

of Aerospace Medicine at Wright-Patterson Air Force 

Base, OH, through the Epidemiology Consult Service. 

Entomology services include the capability to identify 

both wide range of arthropods from around the world 

and pathogen surveillance including the detection of 

several arboviruses, Bartonella spp., Leishmania spp., 

Plasmodium spp., and Rickettsia spp.5-10 Many of these 

identifications and pathogen screening services are used 

for both threat assessments and as part of standard pest 

management operations.

Long-term data on both vectors and associated pathogens are maintained for later review and determination 

of epidemiologic patterns. With the ongoing conflicts in 

the Middle East, analysis of trends in pathogen surveillance continues to be relevant. Several mosquito-borne 

arboviruses including West Nile virus (WNV) and 

Sindbis virus (SINV) were historically reported from 

both military members and local mosquitoes in the region.11,12 We reviewed the arbovirus surveillance data for 

Incirlik Air Base (AB) in Turkey from 2011-2016 with a 

discussion of 2 major viruses.

Incirlik AB is in a strategically relevant location. It is 

located in far southern Turkey relatively near Syria and 

other Middle Eastern countries, including Iraq, Lebanon, and Israel. Surveillance is relevant for ongoing 

force health protection. In addition, pathogens detected 

in Incirlik could be useful in understanding viruses 

circulating in neighboring countries. Several mosquitoborne viruses such as West Nile and Sindbis are endemic in Turkey.13,14

Methods

The US Air Force currently maintains several military 

bases in the Middle East, but Incirlik AB, which opened 

in February 1955, has one of the longest established 

records. Most bases conduct vector and pathogen surveillance programs. Mosquitoes on many of these bases breed in water retention ponds or subsurface catch 

basins below parking lots or streets. Mosquitoes were 

trapped on the air base using the solid-state Army miniature traps with dry ice. Traps were set out seasonally 

from April through October and run on a weekly basis. 

Mosquito fogging typically occurred 3 days a week. Two 

traps were used, both near the perimeter fence, one near 

the sewage treatment plant, and one near base quarters. 

Mosquitoes were killed by freezing and shipped dry to 

the USAF School of Aerospace Medicine for identification and pathogen detection. Mosquitoes were morphologically identified using regional keys.15,16 Voucher 

specimens were deposited in the Ohio State University 

Museum of Biological Diversity, Columbus, OH, or the 

Walter Reed Biosystematics Unit, Silver Spring, MD.

After identification, mosquitoes were pooled by species and tested for arboviruses. Pools ranged from 1-25 

mosquitoes depending on submission numbers. Most 

Operational Mosquito and Vector-Borne

 Diseases Surveillance at Incirlik Air Base,

 Turkey

Will K. Reeves, PhD, MS

Myrna M. Miller, DVM, PhD

Orhan Bayik, PHS

Maj Leah Chapman, USAF, MS

Abstract

Arboviruses on Incirlik Air Base, Turkey, pose a threat to military personnel and civilians, but might also be 

relevant for understanding the threats in neighboring conflict zones such as Syria. We reviewed 6 years of mosquito and arbovirus surveillance at Incirlik Air Base. Over 6,000 mosquitoes were identified as Aedes caspius, 

Anopheles claviger, Culex mimeticus, Cx. perexiguus, Cx. pipiens, Cx. sinaiticus, and Culiseta longiareolata. 

Almost all of the mosquitoes (more than 90%) were Cx. perexiguus or Cx. pipiens. Both West Nile virus and 

Sindbis virus were detected in 6 mosquito pools among collections made in 2013, 2014, and 2015.
January – June 2017 87

mosquitoes were tested with a RAMP* WNV test and 

the VecTOR Test Systems (Thousand Oaks, California) Alphavirus panel that detects chikungunya (CHIK), 

equine encephalitis, Mayaro, SINV, Venezuelan equine 

encephalitis, and Western equine encephalitis viruses. 

These rapid assays which had been designed for field 

deployment and rapid screening of mosquitoes 17 were 

among the tools used from 2011-2016 by the USAF 

School of Aerospace Medicine. Both the VecTOR Test 

Systems and RAMP assays were performed in accordance with manufacturers’ guidelines. We occasionally used commercially available inactivated West Nile 

and chikungunya virus antigens as a positive control to 

validate the assay sensitivity as suggested.18 As a further validation, we verified 2 of the positive pools by reverse transcription polymerase chain reaction (RT-PCR). 

West Nile virus positive pools were verified using the 

protocols by Lanciotti et al.19 The PCR products were 

sequenced and compared to sequences in GenBank using the BLAST† program.

There were 6 pools of Culex perexiguus Theobald or Cx. 

pipiens L. that tested positive for arboviruses as shown 

in the Table. There were 4 pools positive for WNV: one 

each on October 2 and 9, 2013; August 29, 2014; and 

September 16, 2015. We validated the October 9, 2013 

results with both the RAMP and VecTOR Test Systems 

WNV assays. The RAMP scores ranged from 150 to the 

maximum limit of 740. Both pools tested by RT-PCR 

were positive with cycle threshold scores of 18.14 and 

21.04 respectively which was less than the viral culture 

positive controls. The 919 base-pair sequence of WNV 

envelope protein was a 100% match to WNV isolate 

Spain/2010/H-1b (GenBank # JF719069).

In addition, 2 pools of combined Cx. perexiguus and Cx. 

pipiens from June 18 and 25, 2014, tested strongly positive for an Alphavirus on the VecTOR Test System assay. 

This assay is designed to detect specific viruses, and 

Sindbis is the detectable virus in the region.13

Comment

We detected West Nile and Sindbis viruses in pools 

of Culex spp. from Incirlik AB. The WNV was in the 

Western Mediterranean WNV subtype in the WNV lineage 1 based on sequence similarity. While often underappreciated, both viruses are a true threat to the military. 

Petersen et al20 estimated that WNV caused hundreds 

of thousands of infections in the United States annually 

from 1999 to 2010. Further, WNV was the leading cause 

of vector-borne disease death in USAF-associated individuals since the 1970s.2

 Sindbis virus infection causes 

severe arthritis, fever, and, in rare cases, encephalitis.21

West Nile virus was detected in a total of 6 collections 

made in 2013, 2014, and 2015. This indicates that the virus circulates at some regularity and could pose a threat 

to both active duty, contractors, civilian personnel, and 

dependents. We separated mosquito collection data between sewage or waste water treatment plants and residential areas and detected infected mosquitoes in both 

areas. This virus requires avian hosts for the natural 

enzootic cycle but mosquitoes can be transovarially infected.22 We could not determine the route of infection 

in mosquitoes from Incirlik AB.

Incirlik AB is close to Syria and other conflict zones 

in the Middle East. Detection of 2 potentially serious 

arbovirus threats on a military base in southern Turkey 

could raise concerns for similar disease threats in neighboring countries. This is especially true in conflict zones 

with little to no infrastructure. Infections by either West 

Nile or Sindbis virus often lead to mild disease but can 

lead to fatal or debilitating infections and might result in 

costly medical evacuations.

In summary, we reviewed 6 years (2011-2016) of WNV 

and SINV detection in mosquitoes from Incirlik AB. 

Culex perexiguus was the most commonly associated 

mosquito with WNV from the base. It is a member of the 

Cx. univittatus Theobald complex and several of these 

have previously been associated with both WNV and 

SINV with evidence of vertical transmission of WNV.23

Our surveillance data suggests that there are repeated 

and predictable threats from WNV and possibly SINV 

on the base.

Acknowledgements

We thank the Public Health and Civil Engineering personnel at Incirlik AB for mosquito surveillance and timely submission of samples. We also thank the current and 

former Entomology staff from the US Air Force School 

of Aerospace Medicine.

This research was partially funded by the Global Emerging Infections Surveillance and Response System.

This article was cleared by the Defense Office of Prepublication and Security Review, Security Review# 

17-S-1077.

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THE ARMY MEDICAL DEPARTMENT JOURNAL

*RAMP (rapid analyte measurement platform) is a registered 

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†Basic Local Alignment Search Tool; National Center for Biotechnology Information, Bethesda, MD.
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Mosquito surveillance data from Incirlik Air Base, Turkey, from 2011-2016 with WNV virus and Sindbis virus screening data.

Locality name Collection Dates Species* and Number Arbovirus Test Results

Housing 2 June -19 August 2011 Culex perexiguus (95)

Culex pipiens (23)

Aedes caspius (1)

Culiseta longiareolata (7)

WNV/Alphavirus Negative

Housing 27 April -14 September 2012 Culex perexiguus (133)

Cules pipiens (388)

Culex mimeticus (7)

Aedes caspius (7)

Culiseta longiareolata (6)

WNV/Alphavirus Negative

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Housing 18 April -20 November 2013 Culex perexiguus (73)

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Aedes caspius (17)

Culiseta longiareolata (7)

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positive for WNV virus.

Sewage Treatment 

Plant

18 April -20 November 2013 Culex perexiguus (821)

Culex pipiens (252)

Culex sinaiticus (301)

Aedes caspius (32)

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positive for WNV virus.

Flight Line 8 April-6 November 2013 Culex pipiens (8) WNV/Alphavirus Negative

Housing 16 May-29 August 2014 Anopheles claviger (1)

Culex perexiguus (658)

Culex pipiens (153)

Culiseta longiareolata (1)

WNV/Alphavirus Negative

Dining Facility 2 July 2014 Culex perexiguus (1)

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Sewage Treatment 

plant

23 April -29 August 2014 Culex perexiguus (327)

Culex pipiens (90)

Aedes capensis (9)

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25 June 2014 tested positive for Alphavirus. One pool of 

25 Culex perexiguus from 29 August 2014 was positive 

for WNV virus.

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Culex pipiens (280)

Aedes caspius (11)

Culiseta longiareolata (14)

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was positive for WNV virus.

Sewage Treatment 

Plant

1 May-16 September 2015 Culex perexiguus (831)

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Aedes caspius (2)

Culiseta longiareolata (3)

WNV/Alphavirus Negative

Housing 12 April -12 August 2016 Culex perexiguus (11)

Culex pipiens (33)

Culiseta longiareolata (3)

WNV/Alphavirus Negative

Sewage Treatment 

Plant

31 March -16 October 2016 Culex perexiguus (225)

Culex pipiens (182)

Aedes capensis (3)

WNV/Alphavirus Negative

*Scientific species identification:

Aedes capensis Edwards Culex perexiguus Theobald

Aedes caspius (Pallas) Cules pipiens L.

Culiseta longiareolata Macquart Culex sinaiticus Kirkpatrick

Culex mimeticus Noè

OPERATIONAL MOSQUITO AND VECTOR-BORNE DISEASES SURVEILLANCE

AT INCIRLIK AIR BASE, TURKEY
January – June 2017 89

THE ARMY MEDICAL DEPARTMENT JOURNAL

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commercial West Nile virus antigens as positive 

controls in the rapid analyte measurement platform 

West Nile virus assay. J Am Mosquito Cont Assoc. 

2015;31:371-374.

19. Lanciotti, S, Kerst AJ, Nasci RS, et al. Rapid detection of West Nile virus from human clinical 

specimens, field-collected mosquitoes, and avian 

samples by a TaqMan reverse transcriptase-PCR 

assay. J Clin Microbiol. 2000;38:4066-4071.

20. Petersen LR, Carson PJ, Biggerstaff BJ, Custer B, 

Borchardt SM, Busch MP. Estimated cumulative 

incidence of West Nile virus infection in US adults, 

1999-2010. Epidemiol Infect. 2013;141:591-595.

21. Niklasson B, Espmark A, LeDuc JW, Gargan TP, 

Ennis WA, Tesh RB, Main AJ Jr. Association of a 

Sindbis-like virus with Ockelbo disease in Sweden. 

Am J Trop Med Hyg. 1984;33:1212-1217.

22. Anderson JF, Main AJ. 2006. Importance of vertical and horizontal transmission of West Nile virus 

by Culex pipiens in the Northeastern United States. 

J Infect Dis. 2006;194:1577-1579.

23. Miller BR, Nasci RS, Godsey MS, Savage HM, 

Lutwama JJ, Lanciotti RS, Peters CJ. First field 

evidence for natural vertical transmission of West 

Nile virus in Culex univittatus complex mosquitoes from rift valley province, Kenya. Am J Trop

Med Hyg. 2000;62:240-246.

Authors

When this study was conducted, Dr Reeves was the 

USAF consult entomologist at the the USAF School of 

Aerospace Medicine. He currently is a regulatory analyst 

at the US Department of Agriculture Animal and Plant 

Health Inspection Service, Biotechnology Regulatory 

Service, Fort Collins, Colorado.

Dr Miller is the chief virologist at the Wyoming State 

Veterinary Laboratory, Laramie, Wyoming.

Mr Bayik is the Public Health Specialist and Food Safety 

and Medical Entomology Program Manager at the Incirlik Air Base, Turkey.

When this study was conducted, Maj Chapman was a 

Public Health instructor at the USAF School of Aerospace Medicine. She currently is the Assistant Contingency Liaison Officer at the Armed Forces Pest Management Board, Silver Spring, Maryland.
90 http://www.cs.amedd.army.mil/amedd_journal.aspx

Vector-borne diseases (VBDs) are among the leading 

causes of morbidity and mortality worldwide. The history of US military medicine dates from the formation 

of the Continental Army in the 1770s.1

 From the inception of our nation, the US military has combated infectious diseases. Today, scientific research programs at 

Walter Reed Army Institute of Research, the Uniformed 

Services University of the Health Sciences, and other institutions are major components in this effort. Additionally, the armed forces have worked to prevent the spread 

of disease through active public health measures such as 

the establishment of vector control programs at military 

bases, and international scientific collaborations aimed 

at increasing our prevention proficiencies.2

 While VBDs 

are of considerable concern for internationally deployed 

active military and civilian personnel, the concern for 

acquisition of VBDs in US territories and states has also 

been a reality for centuries. Endemic, emerging, and 

newly introduced VBDs have spread across the United 

States and its territories, creating foci of disease transmission. Military personnel are particularly susceptible 

to VBDs due to their increased contact with vectors during outdoor training exercises, military response missions, and occupational-specific exposures.

US Military Entomology Infrastructure

Vector control and pest management across the Department of Defense (DoD) involves a multitude of organizations both internal and external to the DoD. Born from 

a need to address the countless casualties and mission 

cost attributed to VBDs during World War II, the Army 

War Department established the Army Committee for 

Insect and Rodent Control (ACIRC) in 1944. In 1956, 

DoD Instruction 5154.12 established the Armed Forces 

Pest Control Board replacing the ACIRC. By 1979, the 

title was amended to its current form, Armed Forces 

Pest Management Board (AFPMB), to better reflect the 

goal of balancing vector and pest control with environmental protection. The AFPMB is composed of pest 

management and medical entomologists from the Air 

Force, Army, Navy, and the Defense Logistics Agency 

(DLA). It has a strategic mandate from the Office of the 

Assistant Secretary of Defense for Acquisition, Technology and Logistics to recommend policy, provide guidance, and coordinate the exchange of information on 

all matters related to pest management throughout the 

DoD. Service entities including the Army Public Health 

Center, Navy Entomology Center of Excellence, Walter 

Reed Army Institute of Research, Walter Reed Biosystematics Unit, and Armed Forces Health Surveillance 

Center assist in the coordination and implementation of 

DoD and/or service-specific pest management policies 

and guidance. The AFPMB is the lead pest management agency for the DoD, coordinating with other US 

government (USG) agencies, non-USG organizations, 

and other foreign organizations as well, including the 

Environmental Protection Agency, US Department of 

Agriculture, US Centers for Disease Control and Prevention (CDC), North Atlantic Treaty Organization, and 

the World Health Organization.

Vector control programs employed by the US armed 

forces date from the late 1800s, when mosquitoes were 

first identified as vectors for human disease pathogens.3

Vector management programs evolved into their modern 

form during the 1970s with the development of integrated mosquito control programs focused on a multifaceted 

approach designed to manage the target pest with integrated measures to mitigate risk of vector disease transmission while balancing risk of pesticide exposure and 

environmental risk. The introduction of West Nile virus 

into naïve US mosquito populations 4

 and the recent chikungunya-Zika Latin American epidemics have created 

a shift in the paradigm of vector control programs.5

 Programs have now begun to balance the concern for environmental effects of pesticide use on a large scale with 

the threat of invasive mosquito species and their respective capacities to introduce emerging VBDs within our 

borders. Currently, several endemic VBDs continue to 

threaten our military and civilian populations residing 

Vector-Borne Diseases of Public Health

 Importance for Personnel on Military

 Installations in the United States

Melissa N. Garcia, PhD, MPH Walter Roachell, MS

Thomas L. Cropper, DVM, MPVM, ACVPM Shannon E. Ronca, PhD, MPH

Sarah M. Gunter, PhD, MPH Ralph A. Stidham, DHSc, MPH

Mathew M. Kramm, PhD Maj Bryant J. Webber, USAF, MC

Maj Mary T. Pawlak, USAF, MC Lt Col Heather C. Yun, USAF, MC
January – June 2017 91

within US borders. Additionally, emerging arboviral 

diseases in Latin America threaten introduction into 

our resident mosquito populations,6,7 and new pathogens 

are being identified on a regular basis through surveillance and innovative pathogen detection research investigations.8-10 In this article, we present an overview of 

vector-borne diseases acquired by military personnel 

during field exercises and/or training evolutions within US borders and end with diseases of future concern 

and conclusions to reduce disease transmission in this 

population.

Mosquito-borne Diseases

West Nile Virus

West Nile virus (WNV) is transmitted by a variety of 

mosquito species found in the United States, primarily 

Culex pipiens Linneaeus in the north, Cx. quinquefasciatus Say in the south, and Cx. tarsalis Coquillett in the 

western states. Over 2,000 cases of WNV occur annually in the United States11 with cyclic peak annual outbreaks.12 Although many infected humans will not show 

signs of disease, 20% develop febrile illness, and 1% 

develop neuroinvasive disease (acute flaccid paralysis, 

meningitis, encephalitis or meningoencephalitis).13 Neuroinvasive disease is a particularly concerning clinical 

manifestation, as these patients have the highest risk of 

long-term morbidity and death.11 Diagnosis is made by 

detection of WNV viremia, or antibodies in the serum 

or cerebrospinal fluid. Acute disease diagnosis is complicated by the short duration of viremia antecedent to 

symptom onset and the sustained presence of IgM infection several months to years post-onset.14 Evidence for 

persistent infection and/or sequelae exists, with patients 

continuing to report morbidity up to 11 years post-infection.15,16 The economic burden from WNV infection is 

considerable with each individual case of acute neuroinvasive disease resulting in up to $51,240 in healthcare 

costs17 and up to $400,000 in long-term loss of productivity wages.18 Currently, there are no Food and Drug Administration approved treatments or vaccines for WNV 

infection; however, vaccine clinical trials are undergoing.19 Surveillance has identified WNV activity at over 

44 DoD sites,20 resulting in nationwide vector control 

efforts.21 The Armed Forces Health Surveillance Center 

reported 323 confirmed cases of WNV illnesses among 

Army, Navy, Marine Corps, Air Force, and Coast Guard 

personnel from 2006-2015.22 An additional 245 cases 

were reported during that time, but the military branch 

of the patients was unknown and those cases could include contractors, civilians, and foreign nationals.

Dengue Virus

Dengue virus (DENV) is transmitted by Aedes aegypti (L.) and Ae. albopictus Skuse mosquitos found 

throughout the southern United States, with the Ae.

albopictus vector geographic range reaching north to 

Minnesota and Maine.23 Globally, 4 serotypes (DENV 

1-4) have been well characterized, with a fifth serotype 

having recently been proposed in Malaysia.24 Infection 

with one serotype does not provide cross-protection 

from other serotypes, and multiple serotype exposures 

increases one’s odds of developing severe clinical outcomes, such as dengue hemorrhagic fever and shock 

syndrome. Symptoms of DENV fever are nonspecific 

(fever, headache, joint, muscle, and bone pain), with 

symptom onset typically occurring 4-7 days after vector 

transmission and lasting 3-10 days after symptom onset. 

As the fever is residing, warning signs for serious clinical manifestations can present, including capillary leakage, marked temperature change, thrombocytopenia, 

change in mental status, rapid weak pulse, and hemorrhagic manifestations that can rapidly progress to circulatory system failure and shock.25 Although the majority 

of dengue cases in the United States are travel-related, 

autochthonous transmission has occurred along the Texas/Mexico border, as well as in Florida and Hawaii.26-30

Diagnosis is determined by laboratory confirmation of 

viremia or antibodies. There is no specific treatment for 

DENV infection; however, fluid replacement treatment 

and pain relievers may improve outcomes among critically ill patients. Vaccine trials are underway31; but controlling mosquito populations in the interim is the best 

method for disease prevention. Between 2006 and 2015, 

approximately 700 cases of dengue fever were reported 

in military personnel.32 Due to the rarity of the locallyacquired disease in most of the United States, one would 

assume most individuals acquired the infection during 

deployment to endemic regions. While only focal outbreaks of autochthonous transmission have occurred in 

the United States, military installations in regions where 

Ae. albopictus and Ae. aegypti exist should monitor for 

the potential of autochthonous infections.26-30

Other Endemic Arboviral Infections of Concern

St. Louis encephalitis (SLE) is a Culex sp. transmitted 

flavivirus infection of notable historical importance,33 as 

this disease served as justification for the foundation of 

new vector control authorities across the country. While 

SLE is less prevalent due to the recent establishment of 

WNV, cases are still reported annually and it is a regular 

contributor to neuroinvasive disease nationally.34 Alphaviruses are another group of positive-sense RNA viruses transmitted through the bite of an infected mosquito 

and are a member of the Togaviridae family.35 They 

typically result in encephalitis or arthralgia. Western 

equine encephalitis (WEE), Eastern equine encephalitis 

(EEE), and Venezuelan equine encephalitis (VEE) viruses are encephalitic alphaviruses, while chikungunya 

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is an arthralgic alphavirus. Aedes, Culex, Psorohpora,

and Culiseta are all known to transmit alphavirus infections. For the encephalitic alphaviruses, most infections 

are asymptomatic, but febrile illness can onset 2-10 days 

after infection and progress to encephalitis in approximately 5% of people.35 Most EEE cases are reported in 

Florida, Georgia, Massachusetts, and New Jersey, but 

transmission is most common around the freshwater 

hardwood swamps in Atlantic states, Gulf Coast states, 

and the Great Lakes regions.36 Most WEE cases occur in Texas, Colorado, Oklahoma, and New Mexico.35

Most VEE cases occur in Central and South America, 

but spillover cases have occurred in Texas.37 Complications of encephalitis can occur with lifetime care costs 

exceeding $4.6 million per patient.35 According to the 

Armed Forces Health Surveillance Center, at least 107 

cases of mosquito-borne viral encephalitedes (contributable to either WEE, EEE, or other less common arboviruses) were reported between 2005 and 2014, with 

cases originating annually from the Marines, Army, Air 

Force, and Navy,38 further demonstrating their contributory role in serviceman illness.

Tick-borne diseases

Lyme disease

Lyme disease, also referred to as Lyme borreliosis, is 

caused by different genospecies of the bacterium Borrelia burgdorferi Johnson sensu lato.39 This vectorborne infection is endemic to the majority of the northern hemisphere with active transmission ongoing in 

Europe, Asia, and the United States.40 Multiple species 

of the Ixodes ticks are capable of propagating sylvatic 

transmission of this pathogen in the United States, but 

only two are implicated in human transmission: the 

black legged tick (also called the deer tick) (Ixodes scapularis Say), and the western black-legged tick (I. pacificus Cooley and Kohls).41 Initial symptoms after exposure through the bite of an infected tick often include the 

development of a rash around the sight of inoculation 

referred to as erythema migrans, colloquially called a 

“bulls-eye” rash. In addition, many patients concurrently 

experience fatigue, headache, arthralgia, malaise, and 

myalgia. Rarely the disease can disseminate after initial 

infection and cause symptoms including carditis, neurologic complications, and arthritis.42 Oral antibiotics 

such as doxycycline, amoxicillin, or cefuroxime axetil 

are commonly prescribed for treatment of Lyme disease.43 Patients treated rapidly after onset of infection 

often recover completely. However, delayed diagnosis 

and treatment may lead to a higher likelihood of developing severe disease. Lyme disease is among the most 

commonly reported VBDs in the United States, with 

an annual incidence of around 300,000 cases.44 It has 

long been proposed that individuals with occupations or 

hobbies that require extended time spent outdoors, such 

as military training exercises, in endemic areas are at 

elevated risk for contracting Lyme disease.45

Historically, the armed forces have struggled with Lyme 

disease infection in military trainees, active duty Soldiers, military dependents, and civilian contractors 

working on military bases and installations. Reports of 

infection date back to 2 years after the pathogen was 

first isolated. A naval base in New Jersey reported an 

incidence rate of 1,063 cases per 100,000 personnel between 1981 and 1982. Identification was based on clinical diagnosis as serologic testing was not yet available, 

introducing the possibility that this number underrepresents the true burden of disease due to misdiagnosis.46

Further highlighting this issue, 2 case reports identified 

military personnel that were not identified until disseminated Lyme disease had developed, and presented 

with rare symptoms including carditis and neurologic 

complications.47,48 The late 1990s was a period of low 

incidence. It was estimated that only 6 seroconversions 

occurred during military duty per 100,000 persons.49,50

Reports of Lyme disease among all branches of the military steadily increased in the early 2000s.51,52

In 2006, an entire 110-person unit was preemptively 

treated for Lyme disease after a training exercise at Fort 

Dix, New Jersey. Between one and 2 weeks after the exercise, at least 5 personnel were diagnosed with erythema migrans. Watchful waiting was judged to be too high 

of a risk for this unit as they were preparing to deploy to 

an austere location in less than 2 weeks. Given the risk 

of cardiac and neurologic complications presenting in 

such circumstances, the entire unit received a 2-week 

course of doxycycline for early Lyme.53 In 2011, a spike 

in cases was reported, at its peak 16 per 100,000 active duty personnel and 25 per 100,000 reservists were 

screened positive for B. burgdorferi exposure.54 Incidence of military exposure appear to correlate with base 

location, with the highest incidence in the northeastern 

United States.55 Reports indicate that a correlation exists between pathogen quantity in ticks removed from 

military personnel and human prevalence of disease at 

a given base, indicating that this may serve as an effective surveillance tool for detection of high risk areas and 

prevention of disease outbreaks.56 This method might be 

particularly useful to distinguish between transmission 

sources for returning serviceman from European installations where transmission is also possible.57

Ehrlichiosis

First reported in the United States in 1986, human ehrlichiosis is caused by infection with either Ehrlichia

chaffeensis Dumler (human monocyte ehrlichiosis) or E.

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ewingii Dumler.58 This zoonotic pathogen is an obligate 

intracellular bacterium that often infects monocytes, 

forming distinct Ehrlichia colonies. Cases are most often reported in the southcentral and eastern regions of 

the United States. This corresponds to the geographic 

region of naturally occurring lonestar ticks (Amblyomma americanum Linnaeus), the primary vector for both 

Ehrlichia species.59 Within one to 2 weeks of exposure, 

patients typically develop general febrile illness and a 

subset present with a wide range of rash (maculopapular to petechial). If not identified and treated promptly, 

ehrlichiosis can cause more severe symptoms including 

difficulty breathing and abnormal bleeding, with a 1.8% 

case fatality.60 The primary treatment is oral doxycycline 

for both adult and pediatric cases.61 It is recommended 

that the antibiotic be prescribed even in suspected cases 

due to the severe and even fatal nature of this infection.

Among the first reports characterizing the causative 

agent of human ehrlichiosis includes a case-report of an 

Army reservist with tick exposure during a field exercise 

at Fort Chaffee, Arkansas.62 Following this discovery, 

a prospective serologic investigation was launched to 

determine seroprevalence of Ehrlichia at Fort Chaffee 

and surrounding bases. Seroconversion was detected in 

1.3% of the Soldiers with available pre-exposure samples 

(n=1,194) with 33.3% of seropositive personnel reporting a previous clinical history consistent with ehrlichiosis. Additionally, seropositive military personnel were 

significantly more likely to report history of tick attachment (RR=3.56, P<.2), indicating that active tick-borne 

transmission of Ehrlichia was ongoing during field exercises at bases within Arkansas.63 A second outbreak 

was detected at a New Jersey base, where 12% of personnel screened seropositive (n=74) with all 9 cases recalling tick exposure during field exercises in knee-high 

grass.64 Additional sporadic human case reports and Ehrlichia pathogen positive ticks collected from military 

personnel indicate this is an ongoing threat to military 

personnel performing field exercises in the eastern and 

southcentral portions of the United States.52,65,66

Rocky Mountain Spotted Fever

Rocky Mountain spotted fever (RMSF, infectious agent 

Rickettsia rickettsia Ricketts) is endemic to the United 

States,67 and is sometimes referred to as spotted fever 

group rickettsiosis when including other less commonly 

transmitted rickettsial pathogens (R. parkeri Lackman 

and Rickettsia spp. 364D). Transmission has been noted 

throughout the contiguous 48 states, with a burden of 

disease occurring in the southcentral states. The primary vector for RMSF is the American dog tick (Dermacentor variablilis (Say)).68 Symptoms of disease present 

2 to 14 days after exposure and typically begin with a 

nonspecific fever and headache. Infection is often misdiagnosed at onset, due to the initial nonspecific nature 

of disease. Typically, it is not until the characteristic rash 

associated with infection forms 2 to 5 days following 

onset of symptoms, that the disease is positively identified.68 Some individuals will have a more severe infection that can lead to vasculitis, and abnormal bleeding 

in the brain and/or vital organs. Those who experience 

severe disease symptoms often suffer from sequelae as 

a result of infection. Diagnosis of RMSF can be difficult 

as no serologic test are available to detect acute infection, and is largely determined by clinical symptoms 

and epidemiologic exposures. Once a case is identified, 

prompt doxycycline administration is critical to limit 

disease severity.67 Given the frequency of movements 

of military trainees from one geographic location to 

another, this often presents well outside its usual geographic range. A thorough travel history is an essential 

component for the patient exam of any military personnel presenting with an acute febrile illness in order to 

consider geographically limited but life-threatening infections like RMSF.

Rocky Mountain spotted fever has affected military 

personnel and working dogs stationed across the United States since 1982. A serosurvey of dogs (N=467) 

housed at 4 different military bases across the country 

found 32% were seropositive, with a range of 4.3% to 

63.4% depending on region. Additionally, a higher rate 

of RMSF exposure was reported (87%) in the working/

sporting dogs breeds screened.69 Prevalence in canine 

populations were seemingly in parallel with human seroprevalence in the same geographic region during this 

time period, suggesting that dogs may provide an efficient sentinel for infection.69,70 At the same Arkansas 

base where Yevich and colleagues63 reported Ehrlichia infections, a serosurveillance study detected 2.5% 

of military personnel (n=1,194) had seroconverted for 

RMSF. While less than a quarter of individuals that developed antibodies for RMSF had clinical symptoms 

(8/30), disease status was strongly associated with history of tick bite (RR=4.3 P<.001).63 Reports continue 

to emerge linking cases of RMSF to field exercises in 

the United States,71 detecting seroconversion in military 

troops,52,72 and identification of pathogen from ticks implicated in human bites.65

Tick-borne Relapsing Fever

Tick-borne relapsing fever (TBRF) occurs when Borrelia spirochetes (predominantly B. hermsii Steinhaus 

and B. turicatae Steinhaus) are transmitted to humans 

by Ornithodoros ticks. These soft-shelled Argasidae 

ticks differ from hard Ixodidae ticks in several key characteristics: they have multiple nymphal stages; they feed 
94 http://www.cs.amedd.army.mil/amedd_journal.aspx

rapidly, typically between 15 and 90 minutes; as adults, 

they can feed and reproduce repeatedly, are capable of 

surviving for several years between blood meals73; and 

the spirochetes may colonize their salivary glands, rather than the midgut, allowing for rapid deposition after 

host attachment.74 An infected human usually displays 

an influenza-like illness, often with some degree of altered mental status, after a mean incubation period of 

7 days (range: 4-18 days). Although severe disease is 

atypical, acute respiratory distress syndrome and other 

serious sequelae have been reported. More commonly, 

the initial illness resolves in 3 days (range: 0.5-17 days), 

followed by an afebrile interval of approximately one 

week and then a relapse of fever. Since borreliae can 

vary their surface protein antigens repetitively, multiple 

relapses are possible. Febrile episodes typically become 

shorter and less severe over time. The mortality rate 

is well below 5%, with some fatalities attributed to a 

Jarisch-Herxheimer reaction after antibiotic initiation.73

Diagnosis of TBRF may be confirmed by visualization 

of spirochetes on thick or thin smears using Giemsa or 

Wright stains during febrile episodes, although serologic and molecular techniques are becoming increasingly 

available.75 The mainstay therapy for infected adults is 

a 7-10 day course of oral doxycycline (100 mg every 

12 hours); other oral and parenteral antibiotics are also 

effective.73

Isolated cases or small clusters of TBRF are possible 

during military field training exercises in endemic regions of the country, most notably Florida, Texas, and 

the Pacific West. Sleeping on floors in close proximity 

to the natural habitat of Ornithodoros ticks, such as in 

limestone caves or rodent-infested cabins, is particularly 

risky.76 In the summer of 2015, an Army Soldier contracted TBRF during a 30-day field exercise in northwestern Texas, likely while sleeping in an abandoned 

barn-like structure. He was hospitalized with fever and 

marked thrombocytopenia but recovered rapidly after 

initiation of doxycycline. Postexposure prophylaxis was 

provided to 10 soldiers in his detachment, none of whom 

became ill.77 This case provided the first human isolate 

of B. turicatae 78 and should remind military public 

health personnel to emphasize tick control and personal 

protective measures during field exercises in TBRF endemic areas. When these measures cannot be followed, 

or when the risk remains high despite their implementation, postexposure prophylaxis with oral doxycycline 

should be considered.79

Other Tick-borne Diseases of Concern

Several additional endemic tick-borne diseases exist in 

the United States. Anaplasmosis (Anaplasma phagocytophilum Dumler rickettsial infection) is transmitted 

to humans by I. scapularis ticks and results in approximately 2,600 incident cases annually, primarily in 

northeastern and midwestern states.80 A recent seroprevalence study of the DoD Serum Repository identified 

a 0.11%-2.6% A. phagocytophilium positive rate among 

a geographically diverse sample of servicemen, identifying this pathogen as an important cause of illness 

in military populations.72 Colorado tick fever is another 

cause of undifferentiated febrile illness, occurring primarily at altitudes of 4,000 ft to 10,000 ft and transmitted by D. andersoni Stiles.81,82 It can result in serious 

complications, including pericarditis, meningitis, and/or 

encephalitis. Tularemia infection (caused by Francisella

tularensis Dorofeev) is a potentially life threatening illness that can be transmitted by Dermacentor sp., Amblyomma sp., and Ixodes sp. ticks, among other nonvector 

transmission sources. Tularemia has been reported from 

all contiguous states, but is most common in southcentral states and the Pacific Northwest.83 Powassan encephalitis is a rare but serious viral tick-borne disease 

common in the northeastern United States, primarily 

transmitted by I. cookei Packard vector, although other 

Ixodes sp. and D. andersoni can be vectors.81,82

Other vector-borne diseases

Chagas Disease

An estimated 6 to 8 million people are infected with 

Chagas disease.84 The disease results from infection 

with the protozoan parasite Trypanosoma cruzi Chagas 

is transmitted to humans through one of several routes: 

vector-borne, congenital, blood-borne, oral, and organ 

transmission. It is most commonly acquired via vectorborne transmission.84 More than 130 triatomine species 

in the Americas can be infected by and transmit T. cruzi.

85 In the United States, the greatest diversity is found 

in southwestern states; Triatoma sanguisuga LeConte 

is the most widely distributed, yet T. gerstaeckeri Gerstaecker is the most commonly collected.85 Autochthonous infection potential increases as triatomine species 

adapt to human dwellings, and as human living environments and military field activities expand into areas of 

sylvatic disease.86

Infection occurs when a T. cruzi positive triatomine 

feeds and excreta contaminates the bite wound or mucosal tissue, entering the bloodstream. After an incubation period of 1 to 2 weeks, an acute phase of 8 to 12 

weeks follows. During the acute phase, patients may be 

asymptomatic, have mild symptoms, or local inflammation at the bite site; however, less than 1% will have 

severe acute disease that manifests as acute myocarditis 

and/or pericardial effusion.87 Chronic infection occurs 

in 3 forms: indeterminate, cardiac, or gastrointestinal 

disease. Most infections remain in an indeterminate 

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phase for life and have positive anti-T. cruzi serology, 

but no clinical signs or symptoms.86 Approximately 

20% to 30% of indeterminate cases progress to cardiac 

and/or gastrointestinal disease years or decades later. 

Cardiac disease is detected by abnormal electrocardiogram, and symptomatic disease may present as aneurysm, thrombus formation, or congestive heart failure.87

Less commonly, progression leads to gastrointestinal 

disease affecting the esophagus and/or colon, leading 

to motility disorders, megaesophagus, or megacolon.88

Treatment with nifurtimox or benznidazole may be 

indicated for acute and indeterminate chronic disease 

to decrease symptoms and clinical course, but must be 

obtained from the CDC and administered under an investigational protocol.88 Two case reports of servicemen 

with military and childhood exposures have been published, highlighting the potential for disease transmission among military personnel in the United States.89,90

Other Endemic Vector-borne

Diseases of Concern

Leishmaniasis is a parasitic infection (20+ Leishmania

sp.) transmitted by sand fly vectors (30+ Phlebotomus

sp.), and can manifest clinically as either cutaneous or 

visceral forms of disease. Two genetic lineages exist that 

correspond to either Old World infections (Asia, Middle 

East, and Africa) or New World infections (Western 

Hemisphere). While vector-borne transmission is most 

common, anthroponotic transmission has occurred,91

heightening concern of this disease among our armed 

forces personnel living in close quarters. While military 

cases have not been reported, autochthonous leishmaniasis human cases have been reported from Oklahoma 

and Texas,92 implicating these states as possible transmission risk areas for military operations and training. 

Trench fever (Bartonella quintana Schmincke infection) is transmitted by the human body louse (Pediculus

humanus humanus Linnaeus), and manifests clinically 

as nonspecific febrile illness, bacillary angiomatosis, 

or endocarditis.93 While trench fever was historically a 

major concern in World War I, it has yet to be reported 

among contemporary service personnel; however, high 

rates of infected body louse among American homeless populations 94 indicates a potential risk to military 

personnel in contact with these populations, ie, those 

serving in natural disaster response operations. Murine 

typhus is a typhus group rickettsiosis transmitted by 

rat fleas (Xenopsylla cheopis Rothschild) and cat fleas 

(Ctenocephalides felis Bouché), which can present as 

nonspecific febrile illness, conjunctivitis, and/or heptosplenomegaly. Murine typhus is a reemerging disease 

with geographic distribution in Hawaii,95,96 Texas,97 and 

California.98

Diseases of Future Concern

Should the disorder infect the Army, in the natural way, 

and rage with its usual virulence, we should have more 

to dread from it, than from the Sword of the Enemy.

Letter from General George Washington

to the Continental Army Surgeon General,

Dr William Shippen, February 6, 1777.99

Whether engaging in stateside-based training activities, 

traditional major combat operations, or humanitarian 

assistance missions, US armed forces military personnel have always had to grapple with ubiquitous vectorborne diseases which have rivaled bayonets, bullets, 

missiles, and mortars throughout history as the causes 

of morbidity, mortality, disability, and diminished operational effectiveness. While certain diseases have lost 

their military importance (yellow fever,100 plague,101

and epidemic typhus,102,103), others remain of concern 

(dengue fever,102,104 leishmaniasis,102,105-107 West Nile encephalitis,108 and malaria100,109), and emerging diseases 

have recently occurred (Zika virus110 and chikungunya 

fever 111) that affect operational forces.

Leishmaniasis, characterized by the CDC as a “neglected tropical disease,” persists as a pestilence of future 

concern. The facts that the incidence of cutaneous leishmaniasis among US military during operations in Iraq 

and Afghanistan was substantial,112 and it is endemic 

in 88 countries113 are reasons to keep this condition on 

military preventive medicine’s radar. Notably, there has 

also been an upsurge of cutaneous leishmaniasis among 

Syrian refugees in traditionally nonendemic locations,114

including Europe.113,115 Furthermore, leishmaniasis has 

been identified in Texas and other areas of the southern United States,116,117 complicating the ability to accurately identify origin of infection in returning military 

personnel.

Mayaro virus, of the genus Alphavirus in the family Togaviridae, is a close relative of chikungunya that produces an analogous debilitating arthralgic disease in 

South America. Mayaro could be endemic in regions 

across the continent but camouflaged by the unspecific 

symptoms it shares with other mosquito-borne viruses. 

First isolated from febrile forest workers in Trinidad 

in 1954,118 the etiologic agent of Mayaro fever has been 

identified in French Guiana, Suriname, Venezuela, Peru, 

Bolivia, Brazil,119 and Haiti.120 The apparent primary 

vectors, Haemagogus mosquitoes, inhabit rural settings, a reason that may justify the relative paucity of 

cases and inhibited endemicity. Conversely, Ae. aegypti

mosquitoes have been shown to be competent vectors 

of Mayaro virus,121 signifying that an urban-dwelling 

arthropod could potentially be a vector of this virus on 
96 http://www.cs.amedd.army.mil/amedd_journal.aspx

a global scale. Because Mayaro virus symptoms can resemble those of both chikungunya and dengue, it may 

be underdiagnosed. While the Mayaro virus has not 

been linked with fatal human disease like dengue, primary infections are often more debilitating, with loss 

of productivity for weeks or even months due to severe 

arthralgia.

Military preventive medicine personnel should also be 

on guard for the Oropouche virus,122 the Amazonian 

cousin of the Mayaro virus, which is spread through 

Culex sp. mosquitos and Culicoides paraensis Goeldi 

midges.123 Both vectors are known to have a broader 

distribution124 than the Aedes sp. mosquitoes that carry 

Zika virus.125 Lastly, Ross River virus, which was previously thought to be indigenous to Australia and Papua 

New Guinea by sustaining itself in marsupials,126 has 

been documented in Pacific Island travelers. The spread 

of these new geographic regions suggest the potential 

for further geographic expansion and global epidemic 

potential. Collectively, these arboviral pathogens not 

only infect people via enzootic spillover, but they use 

humans as amplification hosts and represent a tremendous risk for urbanization.127 The latest epidemic activity of Zika and chikungunya should underscore the 

need to consider these diseases in febrile US service 

members returning from endemic areas and serve as a 

caution that presumably obscure viruses like Mayaro 

virus, Oropouche virus, and Ross River virus should 

not be underestimated as potentially emerging human 

pathogens.

Conclusion

Vector-borne diseases have been an important cause of 

morbidity and mortality since the inception of our nation. In line with the general population, military and 

civilian personnel acquire diseases while on stateside 

military installations, active missions, and training exercises. Military personnel with occupational duties resulting in extended time outdoors are potentially at an 

increased risk for VBD transmission. Development and 

implementation of integrated vector management plans 

can be useful tools to reduce vector exposure and transmission risk during disease outbreaks, in endemic disease areas, or in the event of emerging VBDs. As identified by the Armed Forces Health Surveillance Center, 

our military personnel routinely acquire a wide range 

of VBDs spread through mosquitoes, ticks, triatomines, 

and other arthropods. Our current review identified the 

VBDs of greatest public health concern to serviceman 

taking part in military missions and training in the United States. Vector surveillance, insecticide applications, 

and personal protection measures are warranted to prevent future infections.

Acknowledgement

We thank LTC Brian Evans and COL Jaime Blow for 

their contribution to the Armed Forces Pest Management 

Board information in this article.

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Haiti, 2015. Emerg Infect Dis. 2016;22:2000-2002.

121. Long KC, Ziegler SA, Thangamani S, Hausser NL, 

Kochel TJ, Higgs S, Tesh RB. Experimental transmission of Mayaro virus by Aedes aegypti. Am J

Trop Med Hyg. 2011;85:750-757.

122. Vasconcelos HB, Nunes MR, Casseb LM et al. Molecular epidemiology of Oropouche virus, Brazil. 

Emerg Infect Dis. 2011;17:800-806.

123. Pinheiro FP, Travassos da Rosa AP, Gomes ML, 

LeDuc JW, Hoch AL. Transmission of Oropouche 

virus from man to hamster by the midge Culicoides

paraensis. Science. 1982;215:1251-1253.

124. Weaver SC, Reisen WK. Present and future arboviral threats. Antiviral Res. 2010;85:328-345.

125. Musso D, Gubler DJ. Zika virus. Clin Microbiol

Rev. 2016;29:487-524.

126. Klapsing P, MacLean JD, Glaze S, McClean KL, 

Drebot MA, Lanciotti RS, Campbell GL. Ross 

River virus disease reemergence, Fiji, 2003-2004. 

Emerg Infect Dis. 2005;11:613-615.

127. Weaver SC. Urbanization and geographic expansion of zoonotic arboviral diseases: mechanisms 

and potential strategies for prevention. Trends Microbiol. 2013;21:360-363.

Authors

Dr Garcia is an Instructor at the National School of 

Tropical Medicine, Baylor College of Medicine, Houston, 

Texas.

Dr Cropper is Director, Trainee Health Surveillance, 

Joint Base San Antonio-Lackland, Texas.

Dr Gunter is a Postdoctoral Associate with the National 

School of Tropical Medicine, Baylor College of Medicine, Houston, Texas.

Dr Kramm is the Air Force Installations and Mission 

Support Center Entomologist, Joint Base San AntonioLackland, Texas.

Maj Pawlak is a Preventive Medicine Physician, 559th 

Trainee Health Squadron, Wilford Hall Ambulatory Surgical Center, Joint Base San Antonio-Lackland, Texas.

Mr Roachell is an Entomologist with the US Army Public Health Command-Central, Joint Base San AntonioFort Sam Houston, Texas.

Dr Ronca is a Postdoctoral Associate with the National 

School of Tropical Medicine, Baylor College of Medicine, Houston, Texas.

Dr Stidham is an Epidemiologist with the Epidemiology and Disease Surveillance Division, US Army Public 

Health Command-Central, Joint Base San Antonio-Fort 

Sam Houston, Texas.

Maj Webber is the Preventive Medicine Element Chief, 

559th Trainee Health Squadron, Wilford Hall Ambulatory Surgical Center, Joint Base San Antonio-Lackland, 

Texas.

Lt Col Yun is assigned to the Infectious Disease Service, 

San Antonio Military Medical Center, JBSA-Fort Sam 

Houston, Texas, and is the Director of the San Antonio 

Uniformed Services Health Education Consortium Infectious Disease Fellowship Program.
102 http://www.cs.amedd.army.mil/amedd_journal.aspx

Traumatic Brain Injury in the United States

Traumatic brain injury (TBI) is a disruption in brain 

function or pathology due to an external force that exceeds the protective capacity of the head and causes 

neuropsychiatric impairment, either permanent or temporary. Complications of TBI include seizures, spasticity, gait abnormalities, agitation, depression, headaches, 

insomnia, cognitive decline, dementia, gastrointestinal 

and urogenital complications, and chronic traumatic encephalopathy. In the United States alone, an estimated 1.7 

million people suffer a TBI annually, of which 275,000 

require hospitalization.1 Falls and motor vehicle accidents are the major causes of TBI in the United States.1

Traumatic brain injury is a leading cause of death and 

disability for individuals between the ages of one and 

44 years, and it is estimated that almost 2% of the population currently live with complications and disabilities 

related to a prior TBI.2

Traumatic Brain Injury in the US Military

Traumatic brain injury is a major health concern for the 

US military. With the widespread use by enemy combatants of improvised explosive devices in recent conflicts, 

there have been a higher proportion of explosive-related 

head injuries in comparison to previous wars.3,4 Data 

from Operation Enduring Freedom, Operation Iraqi 

Freedom, and Operation New Dawn show that blastrelated injuries are a major cause of TBI for military 

troops.5

 As illustrated in the Figure, from 2000-2016, 

there were 357,048 medical diagnoses of traumatic brain 

injuries in the US military with 82.3% of these injuries 

classified as mild TBIs.6

 A mild TBI is characterized by 

the Department of Defense as a confused or disoriented 

state that lasts less than 24 hours; or loss of consciousness for up to 30 minutes; or memory loss lasting less 

than 24 hours. Mild TBI is difficult to detect as there is 

often a lack of any external evidence of damage to the 

head. A CT scan is generally not indicated for these patients, and if obtained, is often found to be normal.6

 Currently, there are no specific laboratory markers or imaging studies that are routinely performed that can detect 

any long-lasting damage to these patients.7

 Due to the 

difficult nature in diagnosing a TBI, there are likely far 

more mild TBIs that remain undiagnosed in the military.

Traumatic Brain Injury and a History of

Neuropsychiatric Dysfunction in War: Shell

Shock, Posttraumatic Stress Disorder, and

Dementia

The development of neuropsychiatric symptoms in military veterans exposed to high-intensity explosives has 

been recognized since World War I (WWI). Military 

troops involved in trench warfare were often barraged 

with artillery, with many Soldiers developing the neuropsychological impairment known as “shell shock.” 8,9

Symptoms of shell shock included fatigue, confusion, 

nightmares and impaired sight or hearing. Soldiers were 

often diagnosed once they became unable to function/

perform their duties, usually without any obvious cause 

that could be identified. Since Soldiers often lacked 

physical signs of disease, they were often accused of 

cowardice or malingering. It was later recognized that 

Blast-Associated Traumatic Brain Injury in the

 Military as a Potential Trigger for Dementia

 and Chronic Traumatic Encephalopathy

Jamal Hasoon, MD

Abstract

Traumatic brain injuries (TBIs) resulting from blast exposures have significantly affected US military personnel throughout the world, particularly in Iraq and Afghanistan. From 2000-2016, more than 350,000 military 

service members were diagnosed with a TBI. Since the majority of blast-related TBIs are classified as mild 

with no definitive findings on physical examination or diagnostic studies, it is difficult to accurately diagnose 

individuals inflicted with such injuries. There are likely far more mild TBIs that remain undiagnosed in the 

military. Traumatic brain injury is a serious public health concern since it can lead to long-term neuropsychiatric changes such as posttraumatic stress disorder and various forms of dementia. Blast-related TBI has also 

been linked with neuropsychiatric dysfunction that is commonly seen in athletes that have suffered chronic 

traumatic encephalopathy. Further research is needed to aid in prevention, diagnostic studies, and care of military service members and veterans who have suffered a TBI.
January – June 2017 103

shell shock shared similar symptoms to posttraumatic 

stress disorder (PTSD), which would later be recognized during the Vietnam War.

Since the conflicts in Vietnam, Iraq, and Afghanistan, 

there has been more insight into the neuropsychological dysfunction of military personnel who suffer from 

PTSD. Similar to the shell shock observed in WWI, 

PTSD symptoms include disturbing thoughts, flashbacks during trauma-related cues, cognitive and behavioral changes, depression, anxiety, and a higher risk of 

suicide. Although there is tremendous overlap between 

PTSD and shell shock in several ways, it is important to 

acknowledge that patients can also develop PTSD without evidence of TBI or head trauma. Examples include 

victims of rape, natural disaster survivors, or individuals 

who have suffered severe psychological stressors. However, evidence shows a TBI alone can be a potential trigger for PTSD as well. A survey of 2,525 US Army Infantry Soldiers deployed to Iraq for one year demonstrated 

that 44% of Soldiers who suffered a mild TBI with subsequent loss of consciousness met the criteria for PTSD.10

Another study found blast-related injuries on mice under 

anesthesia produced PTSD behavioral changes in the 

absence of psychological stressors.11 Traumatic brain 

injury appears to be a very strong risk factor for the 

development of PTSD. It is also very likely that the majority of Soldiers who suffered from shell shock in WWI 

would fit the diagnosis of PTSD in modern times.

Traumatic brain injury has also been recognized as a potential risk factor for the development of dementia and 

neurodegeneration.12 Individuals who suffer from dementia typically demonstrate a decline in cognitive ability that is often severe enough to interfere with daily life. 

These individuals may demonstrate memory loss, cognitive decline, and behavioral or psychological changes. 

A study of US Navy and Marine Corps veterans hospitalized during WWII demonstrated an association between TBI in early to midlife with the development of 

dementia later in life.13 Veterans who sustained a TBI 

that resulted in loss of consciousness or posttraumatic 

amnesia longer than 30 minutes but less than 24 hours 

were more than twice as likely to develop dementia.13

Veterans who sustained a TBI severe enough to cause 

loss of consciousness or posttraumatic amnesia greater 

than 24 hours were more than 4 times likely to develop 

dementia compared to the control counterparts.13 These 

findings demonstrate that TBI-related complications 

may manifest much later in life, and our veterans may 

actually have a higher risk of developing dementia compared to the civilian population.

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Traumatic Brain Injury and Chronic Traumatic

Encephalopathy

Chronic traumatic encephalopathy (CTE) is a neurodegenerative disease that is thought to develop secondary 

to repetitive head trauma, including concussive and subconcussive injuries. The clinical presentation of CTE 

varies, and often overlaps with other neuropsychological disorders including Alzheimer’s, Parkinson’s, and 

PTSD. Patients with CTE often exhibit both cognitive 

and behavioral impairment including chronic headaches, memory difficultly, poor impulse control, aggression, depression, suicidal tendencies, and also dementia.14,15 Behavioral changes are typically seen earlier in 

the disease, and these patients may often be diagnosed 

with PTSD. Later in the disease process, patients show 

symptoms such as memory loss and cognitive decline 

consistent with dementia.

Chronic traumatic encephalopathy is a newly recognized disease with few confirmed diagnoses. The disease is a clinical diagnosis that can only be confirmed 

postmortem. It is often regarded as a disease that develops from repetitive head trauma in athletes, such 

as football players and wrestlers, who suffer multiple 

concussive injuries. However, new data suggests military personnel may be at risk for developing CTE, and 

a single blast-related TBI may be sufficient to trigger 

the neuropathologic changes seen in CTE.16 Brain pathology of postmortem athletes who have suffered repetitive concussive injuries have demonstrated distinct 

neuropathologic changes that differentiate CTE from 

other known forms of dementia.15-17 Comparison of 

brains from postmortem military veterans exposed to 

blast injuries demonstrated neuropathology that was 

indistinguishable from the neuropathology of young 

athletes with CTE.16 Furthermore, experiments on rodents exposed to an isolated blast-related injury demonstrated neuropathology similar to that of patients found 

to have CTE from repetitive head trauma.16 This study 

in rodents also showed that TBI from blast-related head 

injuries resulted in progressive neurodegeneration that 

continued for more than one year after the initial injury. 

These studies demonstrate that isolated blast-associated 

TBIs could trigger the development of CTE-like symptoms and neuropathology.16

Comment

Traumatic brain injury is a serious public health concern in both civilian life and in the US military. Today, 

our military troops have a greater risk of TBIs due to 

the increased use of improvised explosive devices and 

subsequent blast-related injuries. Evidence shows that a 

mild TBI is a risk factor for developing long-term neuropsychiatric conditions such as PTSD, dementia, and 

even CTE. Since CTE is a newly discovered disease that 

can currently only be confirmed postmortem, CTE may 

be overlooked as a differential diagnosis and mistaken 

for other neuropsychiatric diseases. Chronic traumatic 

encephalopathy should be considered in patients that 

have a previous history of a TBI and are being evaluated for a neuropsychiatric disorder. Furthermore, the 

diagnosis of CTE may better explain the development of 

symptoms in veterans found to have shell shock, PTSD, 

or other forms of dementia. Any patient with a history 

of TBI should be counseled and closely followed to ensure appropriate care and social support. With increasing awareness that military service members are at risk 

for TBI-related neurodegenerative diseases, further research into diagnostic studies, preventive strategies, and 

eventually therapeutic interventions is essential for the 

wellbeing of our veterans.

References

1. Faul M, Xu L, Wald MW, Coronado VG. Taumatic

Brain Injury in the United States: Emergency Department Visits, Hospitalization, and Deaths 2002-

2006. Atlanta, GA: Centers for Disease Control 

and Prevention, National Center for Injury Prevention and Control; 2010. Available at: https://www.

cdc.gov/traumaticbraininjury/pdf/blue_book.pdf. 

Accessed April 14, 2017.

2. Cicerone KD, Kalmar K. Persistent post-concussive syndrome: structure of subjective complaints 

after mild traumatic brain injury. The Journal of

Head Trauma Rehabilitation. 1995;10(3):1-18.

3. Owens BD, Kragh JF Jr, Wenke JC, Macaitis J, 

Wade CE, Holcomb JB. Combat wounds in Operation Iraqi Freedom and Operation Enduring Freedom. J Trauma. 2008;64:295-299.

4. United States Government Accountability Office. 

Mild traumatic brain injury screening and evaluation implemented for OEF/OIF Veterans, but challenges remain. February 2008 [internet]. Available at: https://digital.library.unt.edu/ark:/67531/

metadc302992/m1/1/. Accessed April 14, 2017.

5. Wojcik BE, Stein CR, Bagg K, Humphrey RJ, 

Orosco J. Traumatic brain injury hospitalizations 

of U.S. Army soldiers deployed to Afghanistan and 

Iraq. Am J Prev Med. 2010;38(suppl):S108-S116.

6. Defense and Veterans Brain Injury Center. DoD 

Worldwide Numbers for TBI [internet]. 2016. 

Available at: http://dvbic.dcoe.mil/dod-worldwidenumbers-tbi. Accessed April 14, 2017.

7. Mac Donald CL, Johnson AM, Cooper D, et 

al. Detection of blast-related traumatic brain injury in U.S. military personnel. N Engl J Med. 

2011;364(22):2091-2100.

BLAST-ASSOCIATED TRAUMATIC BRAIN INJURY IN THE MILITARY AS A POTENTIAL TRIGGER FOR

DEMENTIA AND CHRONIC TRAUMATIC ENCEPHALOPATHY
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8. Mott FW. The effects of high explosives 

upon the central nervous system, II. Lancet. 

1916;4826:441-449.

9. Mott FW. The effects of high explosives 

upon the central nervous system, III. Lancet. 

1916;4828:545-553.

10. Hoge CW, McGurk D, Thomas JL, Cox AL, Engel CC, Castro CA. Mild traumatic brain injury 

in US Soldiers returning from Iraq. N Engl J Med. 

2008;358:453-463.

11. Elder GA, Dorr NP, De Gasperi R, et al. Blast exposure induces post-traumatic stress disorder-related 

traits in a rat model of mild traumatic brain injury. 

J Neurotrauma. 2012;29:2564-2575.

12. Fleminger S, Oliver DL, Lovestone S, Rabe-Hesketh S, Giora A. Head injury as a risk factor for 

Alzheimer’s disease: the evidence 10 years on; a 

partial replication. J Neurol Neurosurg Psychiatry. 

2003;74:857-862.

13. Plassman BL, Havlik RJ, Steffens DC, et al. Documented head injury in early adulthood and risk of 

Alzheimer’s disease and other dementias. Neurology. 2000;55:1158-1166.

14. Stern RA, Daneshvar DH, Baugh CM, et al. Clinical presentation of neuropathologically-confirmed 

chronic traumatic encephalopathy in athletes. Neurology. 2013;81:1122-1129.

15. Baugh CM, Stamm JM, Riley DO, et al. Chronic traumatic encephalopathy: neurodegeneration following repetitive concussive and subconcussive brain trauma. Brain Imaging Behav. 

2012;6:244-254.

16. Goldstein LE, Fisher AM, Tagge CA, et al. Chronic traumatic encephalopathy in blast-exposed 

military veterans and a blast neurotrauma mouse 

model. Sci Transl Med. 2012;4(134):134ra60. 

Available at: https://doi.org/10.1126/scitransl

med.3003716. Accessed April 14, 2017. DOI: 

10.1126/scitranslmed.3003716.

17. Gavett BE, Stern RA, McKee AC. Chronic traumatic encephalopathy: a potential late effect of 

sport-related concussive and subconcussive head 

trauma. Clin Sports Med. 2011;30(1):179-188.

Author

Dr Hasoon is a resident physician in the Department of 

Anesthesiology at the Baylor College of Medicine, Houston, Texas.
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When service members return to their normal routines 

following the end of a war, some of them develop illnesses. These may be associated with heredity, personal 

risk factors such as smoking, normal aging, or the result 

of their occupational exposures. When someone develops a serious illness, it is common for them to review 

their past to look for a potential cause. This is human 

nature and part of the “why did this happen to me?” assessment. For many, their military deployments are likely to be considered. Deployments represent changes to 

routines such as eating and sleeping and lack of control 

over daily life, and may also include changes in hygiene 

and unpleasant exposures to things such as trash burning; old, abandoned warehouses and industrial sites; and 

sources of radiation. For many, a deployment may be the 

most unusual and noteworthy experience in their memory. Therefore, it is not uncommon for returning service 

members to ask their health care providers whether or 

not the deployment was a factor in their illness. Associations between exposures to burning trash, blowing sand 

(particulate matter), old chemical weapons, and subsequent illnesses have been considered following Operations Iraqi Freedom, Enduring Freedom, and New Dawn. 

Generally, few clear associations have emerged. The 

challenges of practicing environmental epidemiology in 

a deployed military force are identified and reviewed.

Approaches in Environmental Epidemiology

Epidemiological studies may be conducted to assess 

potential relationships between exposures during deployment and health outcomes. Epidemiology is a field 

concerned with the identification of factors associated 

with disease and development of public health measures 

for disease reduction. Environmental epidemiology is 

the study of the effect on human health of physical, biologic, and chemical factors in the external environment, 

broadly conceived.1

 It examines associations between 

exposures and outcomes to determine if an association 

places a segment of the population at increased risk of 

certain outcomes, usually a disease. The magnitude of 

the association measures how much increased risk of an 

outcome can be associated with the exposure. Avoidance 

of the exposure should then result in a proportionally 

lowered risk for the outcome. Thus, one aim of public 

health is to optimize health by limiting hazardous exposures. There are several different types of studies which 

may be conducted, the selection of which depends on 

the study question(s) being addressed, the frequency of 

the exposure(s), and outcome(s) of interest, as well as 

the expected strength of association between them, the 

current state of knowledge related to the topic at hand, 

issues regarding efficiency and validity, and matters of 

practicality and ethical considerations.

Cross-sectional Studies

A cross-sectional study attempts to assess exposure and 

outcome at the same time. It provides a snapshot view 

(cross-section) of disease patterns in the population of 

interest while assessing the degree of presumed or documented exposure.2

 The main difficulty with this type of 

study is that the “snapshot” captures only one moment in 

time, and by looking at exposure and outcome at the same 

time, it is not always possible to say with certainty that 

the exposure preceded the outcome. This is a noteworthy 

limitation of cross-sectional studies due to the fact that in 

order to draw conclusions about exposure-outcome causation, epidemiologists must be able to first establish correct temporal sequencing. Also, because cross-sectional 

studies are a snapshot, cases of illness can be counted 

only if present at the time of the look, which presents 

another disadvantage of this study design: it tends to 

identify prevalent cases of long duration and may miss 

capturing individuals who are lost to follow-up shortly after the outcome is established or acute outcomes 

among individuals who recover quickly. Advantages of 

cross-sectional studies include high generalizability and 

minimal resource requirements, relatively speaking, due 

to the fact that they can be conducted quickly.

An example of a cross-sectional study would be testing 

pulmonary function in service members exposed to particulate matter levels in the desert at the time of exposure. 

The aforementioned limitation would be highlighted if a 

unit demonstrated some impairment in pulmonary function, as it would not be possible to say for certain that it 

was due to in-theater particulate matter exposure. It is 

Challenges of Practicing Environmental

 Epidemiology in the US Military

Coleen Baird, MD, MPH

Jessica Sharkey, MPH

Joel Gaydos, MD, MPH
January – June 2017 107

possible that some unit members had another type of 

exposure prior to deployment that influenced their pulmonary function. Individuals who were so affected that 

they had been evacuated from theater, or who were not 

onsite at the time of the testing would not be included.

Cohort Studies

Cohort studies assemble an exposed group and an unexposed group, follow them forward in time for the occurrence of health effects, and compare the rates in the 2 

groups. This study type is suitable when the at-risk population is well-defined, the disease is common, or the 

exposure is rare or specific. When the at-risk population 

is known and there is a specific or rare exposure, it is 

easy to identify the exposed versus the unexposed. The 

advantages of this method are that accurate measures of 

exposure are possible before the outcome occurs, and 

that multiple outcomes can be monitored. A source of 

bias in these studies is that the knowledge of exposure 

status may alter the follow-up and subsequent diagnosis. 

Other limitations include the fact that many years may 

pass between exposure and outcome, which makes the 

study costly, increases the likelihood of losing people to 

follow-up, and is particularly impractical when looking 

at rare diseases or diseases with long latency periods, 

such as cancer. An unexposed population is used to provide a comparison rate for the health effects of interest. 

However, it is critical that the exposed and unexposed 

populations differ only with respect to exposure status, 

something that is not always easily accomplished. To alleviate some of the limitations, nonconcurrent or retrospective cohort studies are often performed. This type 

of study reconstructs exposed and unexposed groups 

based on exposure in the past. These groups are then 

compared with respect to the rate of the outcomes of 

interest. The chief advantage is that since exposure occurred in the past, sufficient time for the outcome to 

develop may have already passed. The disadvantage occurs when exposure is not well quantified at the time of 

occurrence.3

 For example, to study the effects of burn 

pit exposure on the respiratory system, health outcomes 

for individuals identified as having been at base camps 

with burn pits were compared to individuals who were 

at base camps without burn pits. One key limitation was 

that exposures associated with the burn pits had not 

been well characterized.4

Case-control Studies

The case-control method consists of identifying people 

with the disease of interest (cases), and people without 

the disease (controls). The unknown variable to be ascertained is the type, frequency, and duration of past 

exposure(s). Critical to the validity of a case-control 

study is careful selection of controls, in that they come 

from the same population as the cases and exposed and 

unexposed controls have the same likelihood of being selected for inclusion.5

 This type of study is useful 

when the population at risk is not well-defined, when 

the disease is rare and the exposure common, and although only one disease can be investigated at a time, 

multiple risk factors can be assessed. The major weakness of this method is that exposure is investigated 

after disease status is established. This leads to recall 

bias, where those who are diseased are more likely to 

remember and report exposures than are those in good 

health. Furthermore, accuracy can be particularly challenging when conducting case-control studies due to the 

fact that relevant exposures may have occurred many 

years prior to data collection, which can be difficult to 

ascertain and define correctly. To assess the potential 

association between deployment and the development 

of specific cancers, one approach would be to compare 

cancer cases with controls who have no such diagnosis. 

The main variable to be assessed would be whether or 

not the cases had a stronger history of deployment.

Factors Which Affect the Success of Studies

Given that these study designs each have strengths and 

weaknesses, other factors affect the ability of any given 

study to accomplish what it sets out to do—support or 

refute an association between an exposure and an outcome. Unfortunately, these limitations are common in 

environmental epidemiology studies. Illustrating this 

point, the US Department of Veterans Affairs asked the 

Institute of Medicine (IOM) to form a committee, its 

main charge being a determination of the association 

between exposure to burn pits during deployment in 

support of Operations Enduring and Iraqi Freedom and 

subsequent long-term health effects.6 Although the IOM 

task was to respond to a specific exposure, the committee’s conclusions regarding feasibility and design 

issues apply on a much broader scale. They identified 

the major challenges of any epidemiologic study to be 

both exposure assessment and outcome ascertainment. 

Regardless of type of study selected, the committee reiterated the elements characteristic of any well-designed 

epidemiological study, including selection of a relevant 

study population of adequate size, comprehensive assessment of exposure, careful evaluation of health outcomes, reasonable methods for controlling confounding 

and minimizing bias, appropriate statistical analyses, 

and adequate follow-up time.

Magnitude of the Disease Risk

If an exposure is strongly associated with an outcome, 

following a group of people who are exposed and comparing their incidence of illness with that of a group 

without the exposure allows one to calculate the excess 

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risk which can be attributed to the exposure. This is the 

cohort study design. For example, nonsmoking physicians in England were compared with smoking physicians, and followed over time to count the number of 

lung cancers in each group. It became clear that cigarette smoking was associated with a risk of lung cancer, 

perhaps 10 times greater than that seen in nonsmokers.7

In this study design, accurate information about the 

magnitude of exposure is collected for all individuals 

and all individuals should be followed for a period of 

time sufficient to allow the outcome to occur. If an exposure produces a risk which is 4, 5, or even 10 times 

greater than in people without the exposure, it is relatively straightforward to sort this out and quantify the 

risk. In contrast, lung cancer risk from passive smoke 

exposure is thought to be elevated to less than 2 times 

that of nonexposed individuals.8

 This small difference 

is difficult to detect. When an exposure is associated 

with only a slightly elevated risk, that risk sometimes 

“blurs into the baseline” upon investigation. This is due 

to the fact that in order to detect a slightly elevated risk, 

one must be able to distinguish it from the baseline, or 

“usual incidence” of the outcome without any exposure. 

This is the role of control groups. However, small risks 

require that the studies include large numbers of people 

so that the study has sufficient statistical power. Since 

large numbers of service members deployed in the most 

recent conflict, this may not be a problem, but might 

be if the outcome is particularly rare. Typically, the increased risk associated with environmental exposures 

is estimated to be small. It has been estimated that the 

probable range of increased risk for an outcome due to 

chronic, low level exposure is less than 2 times that of 

someone without the exposure.8

 This has implications 

for the number of people necessary in each group in order to detect a difference. This is especially so when 

competing exposures such as smoking may lead to a 

much higher elevation of risk, so the role of smoking 

would have to be thoroughly assessed.

Magnitude of the Exposure

Detection of effects associated with exposure is also 

greatly simplified when the magnitude of the exposure 

is great. Excess numbers of leukemias were easily detected following the exposure to radiation from atomic 

bombs dropped on Hiroshima and Nagasaki because 

the magnitude of exposure was so large.9

 High doses of 

radiation are strongly associated with leukemia as an 

outcome. In contrast, environmental exposures are classically small—doses lower than those known to produce 

effects in animal studies. Often the exposures may be 

chronic low-level exposures, occurring over weeks or 

months, but which may not have been quantified since 

they were not recognized.10 Although some studies ask 

individuals about whether or not they were exposed, they 

do not always know. This can lead to misclassification 

bias, where it becomes difficult to categorize individuals as exposed or nonexposed, and thus difficult to make 

causal interpretations about their outcomes. One study 

evaluated health outcomes for service members who 

were present at the time of a sulfur mine fire that burned 

for several weeks in Iraq.11 The “exposed” group included individuals within a 50 km radius of the site, based on 

satellite images of the plume and rosters of units in the 

area. However, some individuals may not have been present, and it was difficult to assess the level of exposure.

Dose-response Relationship

An axiom of toxicology states, “The dose makes the 

poison.” The implication of this is that the larger the 

dose, the larger the effect. Associations between exposures and outcomes are strengthened when this relationship can be shown to be logical and uniform. This does 

not necessarily mean that the greater the exposure, the 

sicker the individual, but that more outcomes (eg, more 

cases of cancer) occur in the higher exposure groups as 

compared with those less exposed. For example, more 

lung cancers would be expected in groups of individuals who smoke 3 packs of cigarettes a day for 20 years 

versus one pack a day for 5 years. This introduces again 

the requirement for accurate, quantitative exposure data. 

Unfortunately, we can rarely reconstruct individual exposures in deployed settings. For example, the study 

looking at health outcomes associated with having deployed to a base camp with a burn pit was reviewed 

by the IOM for their report, Long Term Health Consequences of Exposure to Burn Pits in Iraq and Afghanistan.4

 One of their criticisms was that the study did not 

identify those with the highest exposures, such as those 

who guarded the burn pits, compared to those with lesser exposures.6

 However, this information on individuals 

was not available. In another example, the study of the 

health outcomes associated with those who were “under 

the plume” of the sulfur mine fire was not able to distinguish individuals who were at locations with higher 

levels of exposure versus those at locations which were 

briefly under the plume.11 This is because the information on exact daily locations of individuals was not 

available. Efforts must be made to accurately measure 

contaminants at the individual level, but having sufficient sampling capabilities at the location of an exposure, particularly an unplanned one, is a daunting task.

Controlling for Confounding

Confounding occurs when an association between exposure and outcome is over or underestimated due to a 

third factor. It is best explained by example. Studies have 

shown that smoking causes an increase in pancreatic 

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January – June 2017 109

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cancer. An investigator attempted to ascertain the relationship between coffee consumption and pancreatic 

cancer. He questioned cases with cancer and controls 

about coffee consumption and found that cases had a 

greater history of exposure to coffee. He concluded that 

coffee consumption was associated with an increased 

risk of pancreatic cancer. Critics challenged his conclusion by pointing out that those who drink coffee are 

more likely to smoke as well, and that by not questioning 

subjects about smoking behaviors and adjusting for it, he 

was in effect seeing the association between cigarettes 

and pancreatic cancer, not coffee and pancreatic cancer.12 Cigarette smoking, the unmeasured “true” risk exposure, was the confounder. In many studies of deployment and health outcomes, individual smoking status is 

unavailable. Furthermore, the rate of smoking has been 

shown to increase on deployments. Since outcomes such 

as respiratory conditions and cancer are strongly associated with smoking, it is an important variable to consider.

Latency Period

The job of the epidemiologist is easier when the time 

from exposure to outcome is short. This time is called 

the “latent period.” An exposure to high levels of chlorine gas leads quite quickly to respiratory distress. Inferring the association between exposure and outcome 

is relatively clear. The same is true in that exposure to 

someone with measles is quickly followed by measles 

in a susceptible person, and ingestion of contaminated 

food quickly leads to gastrointestinal illness in those 

who ingest it. An outbreak or cluster of outcomes is 

clearly defined, and searches for “exposures” and associations ensue.

Unfortunately, in environmental scenarios, most of these 

conditions are not met. Cancer outcomes typically have 

latent periods on the order of decades. This presents 

difficulty in that the exposure can go unrecognized or 

forgotten because it does not lead to an acute outcome; 

and if it eventually leads to an outcome, sufficient time 

may have elapsed that the individual no longer recalls 

the exposure, or that the past “dose” or magnitude of the 

exposure can only be estimated by memory.10 This, as 

mentioned, leads to recall bias.

Specificity of the Health Effect

Phocomelia, the congenital absence of the proximal part 

of a limb, is an uncommon defect in infants. When infants were seen to have this defect in greater than expected numbers, mothers were questioned about the use 

of medications during pregnancy. It became apparent 

that infants with the defect were more likely to be born 

to women who had taken Thalidomide, compared with 

women who had not.13 In this instance, the outcome was 

specific enough to allow for the assembly of a group of 

these mothers to compare with mothers who had normal outcomes, and compare their prenatal exposures. 

Another classic example was when occupational health 

physicians noticed an increased incidence of an unusual 

liver cancer called angiosarcoma in certain workers and 

determined that they had an increased history of exposure to vinyl chloride. The key was that the outcome was 

one with few other causes. Legionnaire’s disease was 

discovered and described because an unusual pneumonia developed with high frequency in a group of men at 

a convention in a single hotel, and was linked to the air 

conditioning system.14 Hantavirus illness in the southwestern United States was characterized after a physician became alarmed that 2 cases of fatal respiratory illness occurred one week apart. Suspicion was raised that 

there was some common exposure.15

When the effect of a low-level hazardous exposure is 

undefined, those exposed may attribute multiple health 

effects from varied causes to the exposure. Individuals 

who have registered for the burn pit registry have reported diverse medical conditions.16 This is likely because they are reporting all health conditions that they 

may have. Without a comparison group, it is difficult 

to determine which effects might be associated with an 

exposure. The lack of a comparison group made it difficult to draw conclusions from clinical evaluations of 

redeployed service members who were veterans of the 

first Gulf War, when there was a concern about potential 

health effects resulting from their exposures in 1991.17

Evaluating the relationship between health effects that 

are known but nonspecific and environmental exposures can also be challenging. Exposures to solvents, 

which are metabolized in the liver, can lead to elevations of liver enzymes in the blood, but so can a host of 

other chemicals including alcohol and acetaminophen 

(confounding). Respiratory symptoms may follow exposure to burn pit smoke, dust storms, or other factors. 

The investigations which attempt to relate exposures to 

effects must take into account other possible exposures 

and question the individuals about these and control for 

these in the data analysis.

Measurement of Health Effect

When studying health effects, it is important to consider 

how the effect is measured. In some studies, health effects were measured as symptoms reported by the study 

participants. Reporting bias may occur when people 

with concerns report their symptoms.1

 Studies which 

use databases that report disease by diagnostic codes are 

subject to problems with miscoding. Analyses of healthcare encounter diagnostic codes following redeployment 
110 http://www.cs.amedd.army.mil/amedd_journal.aspx

are subject to unique limitations. For example, acute 

changes in health status during deployment due to exposures in theater could be missed. Likewise, changes 

in health status that occur over the long-term could be 

delayed beyond the available follow-up period and are 

therefore unobservable. This would be demonstrated, 

and particularly problematic, in evaluations of cancers 

with longer latency periods. Using medical encounter diagnostic codes to define cases may result in false 

positives, capturing as cases those individuals for whom 

health care providers entered codes for conditions that 

were being considered but not yet rejected or confirmed.

Multiple Hypothesis Testing

Since health effects are not always known, a common 

strategy employed in investigations is to question patients about multiple health effects. Questionnaires may 

ask about respiratory symptoms, gastrointestinal symptoms, neurologic symptoms, and vague, nonspecific 

symptoms. This can result in the calculation of a significantly high disease rate due to chance alone. Statistical 

“rules” for study precision generally allow for 5% error 

due to chance. This means that if 100 symptoms were 

asked about, 5 of the 100 could appear to be increased 

due to chance alone.3

Power and Sample Size

The power of a study is its statistical ability to detect 

a difference between 2 groups if it truly exists (for example, an elevated cancer rate) between the exposed and 

the unexposed. The power of a study is intimately related to the sample size, which is the number of people 

in each of the 2 groups. It is also related to the prevalence of the disease of interest in the population. In order to detect significant patterns, rare diseases such as 

cancer in relatively young populations, are studied using 

a case-control approach, or require large populations. 

More common diseases can be studied in smaller populations. However, to the extent that multiple causes (or 

exposures) are involved, as they are with most chronic 

diseases, larger populations are generally required in order to obtain significant results in studies of more common diseases as well. Refining the measures of diseases 

and the assessment of exposure can improve the power 

of a study to detect an association.

Conclusions

Some situations lend themselves to an epidemiologic investigation of the strength of association. The strength 

of an association, while not proof of causation between 

the exposure and the outcome, gives one confidence that 

the two may be related. These situations occur when the 

exposure is large and leads to a greatly increased risk of 

a specific outcome. The exposure must have occurred 

before the outcome, and ideally, the outcome follows 

soon after the exposure. Ideally, current and past exposures to chemicals would be precisely quantified, and 

testable hypotheses of specific health effects would be 

generated. An accurate assessment of adverse health 

effects in exposed and unexposed populations would 

be made, and account would be taken of all potential 

confounders. The sample size would be adequate, and 

irrelevant hypothesis would not be included. Because 

all studies have limitations, these conditions are never 

achieved. However, practitioners of military preventive 

medicine must continue to work to quickly identify potentially hazardous exposures, precisely measure the exposures, and document those who were exposed and the 

extent of their exposures to facilitate these evaluations.

References

1. Schneider D, Lilienfeld DE. Lilienfeld’s Foundations of Epidemiology. 4th ed. New York, NY: Oxford University Press. 2015.

2. Morgenstern H, Thomas D. Principles of study 

design in environmental epidemiology. Environ 

Health Persp. 1993;101(4):23-38.

3. Aschengrau A, Seage, GR: Essentials of Epidemiology in Public Health. London, UK: Jones and 

Bartlett Publishers International; 2003.

4. Abraham JH, Eick-Cost A, Clark LL, et al. A retrospective cohort study of military deployment and 

postdeployment medical encounters for respiratory 

conditions. Mil Med. 2014;179:540-546.

5. Rothman KJ, Greenland S, Lash TL. Case-control 

studies. Encyclopedia of Quantitative Risk Analysis and Assessment. Vol I. West Sussex, England: 

John Wiley & Sons; 2008:192-204.

6. Institute of Medicine. Long Term Health Consequences of Exposure to Burn Pits in Iraq and Afghanistan. Washington DC: The National Academies Press; 2011:1-9,117-128. Available at: https://

www.nap.edu/read/13209/chapter/1. Accessed 

April 6, 2017.

7. Doll R, Hill AB. The mortality of doctors in relation to their smoking habits. BMJ. 

1954;328(7455):1529-1533.

8. Hori M, Tanaka H, Wakai K, Sasazuki S, Katanoda K. Secondhand smoke exposure and risk of lung 

cancer in Japan: a systematic review and metaanalysis of epidemiologic studies. Jpn J Clin Oncol. 

2016;46(10):942-951.

9. National Research Council. Health Risks from Exposure to Low Levels of Ionizing Radiation. BEIR 

VII Phase 2. Washington DC: The National Academies Press; 2006:141-154. Available at: https://

www.nap.edu/read/11340/chapter/1. Accessed 

April 6, 2017.

CHALLENGES OF PRACTICING ENVIRONMENTAL EPIDEMIOLOGY IN THE US MILITARY
January – June 2017 111

THE ARMY MEDICAL DEPARTMENT JOURNAL

10. National Research Council. Environmental Epidemiology, Volume 1: Public Health and Hazardous Wastes. Washington, DC: National Academies 

Press; 1991.

11. Baird CP, DeBakey S, Reid L, Hauschild V, Petruccelli B, Abraham JH. Respiratory health status of 

US Army personnel potentially exposed to smoke 

from 2003 Al-Mishraq Sulfur Plant fire. J Occup 

Environ Med. 2012;54(6):717-723.

12. MacMahon B. Concepts of Multiple Factors. In: 

Lee DHK, Kotin P, eds. Multiple Factors in the 

Causation of Environmentally Induced Disease. 

New York, NY: Academic Press: 1972:chap1.

13. Taussig HB. Thalidomide and phocomelia. Pediatrics 1962;30(4):654-659. Available at: http://pedi

atrics.aappublications.org/content/30/4/654.short. 

Accessed April 6, 2017.

14. Chin J, ed. Control of Communicable Diseases 

Manual. 17th ed. Washington, DC: American Public Health Association; 2000.

15. Centers for Disease Control and Prevention. 

Outbreak of acute illness-southwestern United 

States, 1993. MMWR Morb Mortal Wkly Rep. 

1993;42(22):421-424.

16. National Academies of Sciences, Engineering, and 

Medicine. Assessment of the Department of Veterans Affairs Airborne Hazards and Burn Pit Registry. Washington, DC: The National Academies 

Press; February 28, 2017. Available at: https://www.

nap.edu/catalog/23677/assessment-of-the-depart

ment-of-veterans-affairs-airborne-hazards-andopen-burn-pit-registry. Accessed April 6, 2017.

17. Institute of Medicine. Adequacy of the Comprehensive Clinical Evaluation Program: A Focused 

Assessment. Washington, DC: The National Academies Press; 1997. DOI: https://doi.org10.17226/6004.

Authors

Dr Baird is the Program Manager for the Environmental 

Medicine Program of the Occupational and Environmental Portfolio, US Army Public Health Center, Aberdeen 

Proving Ground, Maryland.

Ms Sharkey is an Epidemiologist for the Environmental 

Medicine Program of the Occupational and Environmental Portfolio, US Army Public Health Center, Aberdeen 

Proving Ground, Maryland.

Dr Gaydos is an Occupational Medicine Physician for 

the Clinical Public Health and Epidemiology Directorate, US Army Public Health Center, Aberdeen Proving 

Ground, Maryland.
112 http://www.cs.amedd.army.mil/amedd_journal.aspx

Historically, lead has been an important commodity for 

industry, including metal alloys, cosmetics, medicinal 

preparations, and paint pigments. Concomitantly, lead 

poisoning has been well described since early Greek 

history (2nd century BC), and, more recently (19th 

century), adverse health outcomes were well defined 

in workers and children.1

 By the 20th century, more indepth studies revealed the cumulative toxic effects of 

lead exposure, especially among the vulnerable population of children.1,2 Today in the United States, there are 

an estimated 4 million households with potential childhood lead exposures, and nearly half a million US children (ages 1-5 years) are known to have blood lead levels 

(BLLs) above 5 μg/dL, the recommended public health 

action level of the Centers for Disease Control and Prevention (CDC).3,4 Although BLLs have demonstrated an 

overall dramatic decline in the past few decades, lead 

exposures among high risk populations (low-income, 

African American, urban, and rural mining communities or developing countries) still exist.5

Interestingly, lead toxicity can affect nearly every body 

system. Because subclinical presentations are common 

among lead exposed patients, this public health risk 

frequently goes unrecognized. Throughout the world, 

children remain a persistent at-risk population due to 

prevalent hand to mouth behavior, close proximity to 

lead exposures, and physiologic higher gastrointestinal 

absorption per unit body weight and increased respiratory rates.5

 Regardless of the route of entry, the toxic 

effects of lead exposures remain uniform.1

 Specifically, 

lead binds to erythrocytes which can then transfer to 

multiple organs (brain, liver, kidneys, spleen, muscles, 

lungs, and heart), and the majority of the lead can absorb 

onto bones and teeth after several weeks.1

 Further, exposures to lead at low levels (BLL less than 10 μg/dL) in 

childhood has also been shown to contribute to deficits 

in central nervous system functioning and cognition.6,7

Children may exhibit “pica,” a unique manifestation 

of elevated BLL which involve abnormal eating habits 

with soil or paint chips.5

Within a child’s environment, main lead exposures 

derive from diet, contaminated soil, paint in homes 

built before 1978, water pumped through leaded pipes, 

imported clay pots, certain consumer products such as 

candies, make-up, jewelry, certain imported home remedies, electronics, and toys.1,3,5 Indirect exposures from 

adult recreational activities such as take-home lead dust 

from firing ranges have also been documented.8,9 More 

importantly, certain parental occupations (ship yards, 

manufacturers, and handlers in lead alloys including 

batteries and ammunition) can also pose considerable 

risk to the child.10 Therefore, occupational lead exposures can indirectly expose the employees’ families and 

children under the age of 6 years by take-home lead dust 

on clothes, boots, hands, and face.3,11-13

In 1991, the CDC responded to this growing public health 

concern by issuing new guidelines.1,3 This included an 

emphasis on the child’s environmental history, parent 

education, and follow-up for children with BLLs of 10 

μg/dL or greater.1

 More recently, the American Academy of Pediatrics emphasized lead screening history in 

a 2005 guidance statement, and the CDC recommended 

initial and follow-up screening (within 1 to 3 months) of 

pregnant and lactating women, neonates, and infants of 

women with BLLs of 5 μg/dL or greater.4 In May 2012, 

the Advisory Committee on Childhood Lead Poisoning 

Prevention recommended the use of a reference range 

for an elevated BLL based on the growing body of literature that levels less than 10 μg/dL do indeed adversely 

affect children.1,3-5,11,14 Thus, the current value (5 μg/dL) 

identifies children with elevated BLLs.3,15 Understanding the complex toxic effects of lead exposures among 

children underscores the importance of primary prevention and vigilant surveillance and screening.

Case Study

An 18-month-old female was seen in the pediatric clinic 

for complaints of “pica.” She was known to eat unusual 

items such as cat food, wood chips from the stair railings, chalk, and paint off the walls. The birth history revealed a normal term delivery via emergency C-section 

with breech presentation without complications or hospitalizations. The mother of the child denied any past 

surgical history, but the child’s past medical history was 

significant for chronic constipation treated with diet and 

medications. Also, her mother expressed concerns of 

child’s development delays in speech, but she denies any 

Indirect Military Occupational Lead Exposure

 to Children at Home: A Case Report

LTC(P) Paul O. Kwon, MC, USA

Acknowledgment: The material in this article has been reviewed by the Walter Reed Army Institute of Research. 

There is no objection to its presentation and/or publication.
January – June 2017 113

neurologic deficits; skin manifestations; bone, teeth, or 

nail deformities. Family history was unremarkable and 

there are no siblings or other family members in the living environment whether foreign or domestic. There was 

no history of travel or visitors from outside the country.

Since September, the family lived on the military installation with well-developed housing (post-1980s) and no 

history of lead in paint, soil, or water systems. The child 

was formula fed with the introduction to whole milk 

at 9 months old, and currently she is solely on whole 

milk with a normal diet of solid foods. Her mother denied any foreign-made toys, ceramics, or products that 

would contain lead. The mother consistently was a “stay 

at home” mom. Her recreational activities included crochet and bow hunting, but she denied any participation 

of the latter since they moved to the current duty station. At the time, The child’s father was a Soldier in the 

US Army; his occupation (Field Artillery) was in small 

arms munitions and Howitzers, but mostly he worked in 

the motor pool. The mother stated that he worked with 

lead products and had coveralls at the workplace; however, he did not change his boots from work to home. 

Although he washed his hands upon entering the home, 

he denied taking any showers prior. According to the 

mother, his boots and duty uniform was typically located in a common place, the laundry room, and were not 

segregated. The child has had full access of the contaminated clothing, and was known to play with the work 

boots constantly. The father’s recreational activities included hunting with a rifle, but he has denied any shooting range activities since arrival at this post. Both father 

and mother drove separate cars, and the child only traveled with the mother in her vehicle. The mother denied 

any history of cross contamination between the car seats.

A review of the medical history showed the 12-month 

visit had normal results on the Ages and Stages Questionnaire (ASQ) and lead screening. Although the subsequent 15-month visit still revealed a negative lead 

screen, the mother reported significant chronic constipation with decreased ASQ scores in fine motor skills. 

All growth parameters were appropriate for age. A follow-up visit with the primary care provider revealed the 

diagnosis of “pica,” although there was still a negative 

lead screening. Laboratory results included a normal 

hematocrit/hemoglobin (12.5/36.8) for her age group. 

However, serum lead levels were elevated (6 μg/dL). 

Public health officials were notified and an evaluation of 

the work and home environment was conducted.

The installation Industrial Hygiene (IH) Department 

conducted a home visit using Lead Check swabs (3M, St 

Paul, MN). Initial qualitative findings revealed positive 

lead contamination only on the father’s uniform and 

boots, with negative results in other high risk areas 

within the home. The IH team then performed a health 

hazard survey of the father’s workplace (Field Artillery 

motor pool) including lead surface sampling. The workers in the motor pool provided routine wheeled vehicle 

and armaments maintenance. Although lead exposure 

can occur both via ingestion and inhalation, airborne 

levels of lead in the motor pool were anticipated to be 

very low risk, whereas the presence of residual lead on 

Field Artillery equipment from firing activities could 

lead to exposure through eating, drinking, and tobacco 

use. One important concern was take-home lead dust on 

uniforms and boots which could contaminate privately 

owned vehicles and home environments. All quantitative samples were collected using Ghost Wipes with 

a 100 cm2

 template. The samples were then sent for 

analysis to the US Army Public Health Command by 

ASTM Method E1613.* The samples were reanalyzed by 

the laboratory using Environmental Protection Agency 

(EPA) Method 200.8† to better quantify lead at the lower 

levels. To note, the IH Group at Brookhaven National 

Laboratory has established industrial guidelines for lead 

dust based on standards developed by the Department of 

Housing and Urban Development and the EPA. Based on 

these guidelines, dust wipe samples standards included 

40 μg/ft2

 for nonlead operational areas. Final sampling 

above the 40 μg/ft2

 cutoff revealed positive for the Soldier’s front of uniform pants, top and bottom of boots, 

and front of the mechanic coveralls, shown in the Table.

Primary exposures to lead were associated with routing maintenance activities. The lead sampling indicated 

the presence of lead in quantities sufficient to warrant 

precautionary procedures to minimize risk of lead exposure to Soldiers and family members. These procedures 

included enforcing the use of dedicated coveralls in the 

workplace and changing areas, not wearing uniform 

boots into the home environment with segregation of 

any potentially contaminated clothing items transported 

THE ARMY MEDICAL DEPARTMENT JOURNAL

*https://www.astm.org/Standards/E1613.htm

†https://www.epa.gov/sites/production/files/2015-08/docu

ments/method_200-8_rev_5-4_1994.pdf

Results of analyses of the home and workplace sampling for 

possible lead contamination.

Item Lead Concentration

(μg/ft2)

Positive 

(>40 μg/ft2)

Front of uniform pants 69.7 Yes

Top and bottom of boots 82.7 Yes

Mechanic desk area <LOQ No

Mechanic work table <LOQ No

Top of mechanic uniform 23.2 No

Mechanic desk area <LOQ No

Front of mechanic coveralls 45.5 Yes

LOQ indicates limit of quantitation.
114 http://www.cs.amedd.army.mil/amedd_journal.aspx

in a plastic bag, prohibited eating, drinking, or using tobacco products in the maintenance section, and integrated washing areas for hands, face, and shower activities 

before interacting with the home environment.

Comment

Indirect occupational exposure to lead in the home environment is a military-relevant topic with enormous 

public health risks. In the mid-1900s, certain US policies 

were enacted to protect children from environmental and 

industrial lead exposures which led to the sharp reduction in children’s BLL (1976 thru 1989).2,16 Thus, legislative actions have proven the effectiveness of public health 

interventions in light of emerging research that elevated 

BLLs in children can cause serious adverse outcomes.17-19

Although the follow-up lead levels, particularly for this 

child, revealed a definitive decline below threshold limits of lead levels after workplace interventions, improvements in early detection, surveillance, and prevention are 

still needed to protect any long term effects to a child’s 

development in the military home. Future studies in 

population risks in the environmental and occupational 

setting among the military are warranted.

References

1. Meyer PA, Brown MJ, Falk H. Global approach to 

reducing lead exposure and poisoning. Mutat Res. 

2008;659:166-175.

2. Levin R, Brown MJ, Kashtock ME, et al. Lead exposures in US children, 2008: implications for prevention. Environ Health Perspect. 2008;116:1285-1293.

3. Centers for Disease Control and Prevention. CDC 

Response to Advisory Committee on Childhood 

Lead Poisoning Prevention Recommendations in 

“Low Level Lead Exposure Harms Children: A Renewed Call for Primary Prevention”. June 7, 2012. 

Available at: http://www.cdc.gov/nceh/lead/AC

CLPP/CDC_Response_Lead_Exposure_Recs.pdf. 

Accessed March 17, 2017.

4. Centers for Disease Control and Prevention. Low 

Level Lead Exposure Harms Children: A Renewed 

Call for Primary Prevention: Report of the Advisory Committee on Childhood Lead Poisoning Prevention of the Centers for Disease Control and Prevention. January 4, 2012. Available: at https://www.

cdc.gov/nceh/lead/acclpp/final_document_030712.

pdf. Accessed March 12, 2017.

5. Ahamed M, Siddiqui MK. Environmental 

lead toxicity and nutritional factors. Clin Nutr. 

2007;26(4):400-408.

6. Needleman HL, Schell A, Bellinger D, Leviton A, 

Allred EN. The long-term effects of exposure to 

low doses of lead in childhood. An 11-year followup report. N Engl J Med. 1990;322:83-88.

7. Surkan PJ, Zhang A, Trachtenberg F, Daniel DB, 

McKinlay S, Bellinger DC. Neuropsychological 

function in children with blood lead levels <10 microg/dL. Neurotoxicology. 2007;28(6):1170-1177.

8. Valway SE, Martyny JW, Miller JR, Cook M, Mangione EJ. Lead absorption in indoor firing range users. Am J Public Health. 1989;79:1029-1032.

9. Bonanno J, Robson MG, Buckley B, Modica M. 

Lead exposure at a covered outdoor firing range. 

Bull Environ Contam Toxicol. 2002;68:315-323.

10. Dolcourt JL, Hamrick HJ, O’Tuama LA, Wooten 

J, Barker EL Jr. Increased lead burden in children 

of battery workers: asymptomatic exposure resulting from contaminated work clothing. Pediatrics. 

1978;62:563-566.

11. Lanphear BP, Hornung R, Ho M, Howard CR, Eberly S, Knauf K. Environmental lead exposure during early childhood. J Pediatr. 2002;140(1):40-47.

12. Lanphear BP, Roghmann KJ. Pathways of 

lead exposure in urban children. Environ Res. 

1997;74:67-73.

13. Charney E, Sayre J, Coulter M. Increased lead absorption in inner-city children: where does the lead 

come from?. Pediatrics. 1980;65:226-231.

14. Gilbert SG, Weiss B. A rationale for lowering the 

blood lead action level from 10 to 2 microg/dL. 

Neurotoxicology. 2006;27(5):693-701.

15. Canfield RL, Henderson CR, Cory-Slechta DA, 

Cox C, Jusko TA, Lanphear BP. Intellectual impairment in children with blood lead concentrations below 10 microg per deciliter. N Engl J Med. 

2003;348(16):1517-1526.

16. Pirkle JL, Kaufmann RB, Brody DJ, Hickman T, 

Gunter E, Paschal DC. Exposure of the US population to lead, 1991-1994. Environ Health Perspect. 

1998;106:745-750.

17. Bellinger DC, Needleman HL. Intellectual Impairment and Blood Lead Levels. N Engl J Med. 

2003;349(5):500-502. Correspondence. Available at: http://www.nejm.org/doi/full/10.1056/

NEJM200307313490515#t=article. Accessed April 

19, 2017.

18. Bellinger DC, Stiles KM, Needleman HL. Lowlevel lead exposure, intelligence and academic 

achievement: a long-term follow-up study. Pediatrics. 1992;90:855-861.

19. Dietrich KN, Ris MD, Succop PA, Berger OG, 

Bornschein RL. Early exposure to lead and juvenile 

delinquency. Neurotoxicol Teratol. 2001;23:511-518.

Author

LTC(P) Kwon is the Director, Preventive Medicine 

Branch, Walter Reed Army Institute of Research, Silver 

Spring, Maryland.

INDIRECT MILITARY OCCUPATIONAL LEAD EXPOSURE TO CHILDREN AT HOME: A CASE REPORT
January – June 2017 115

I. Introduction

The Canadian poet and adventurer Robert W. Service 

could not have imagined the emotional journey he was 

about to embark on when he enlisted in an “American 

Ambulance Unit” in 1915 in Paris. Though unenthusiastic about combat, he did not mind taking “a chance of 

being killed” while helping others. Only a few weeks after enlisting in the ambulance service, the middle-aged 

poet found himself “driving an ambulance under fire” in 

Flanders. Despite the explosions all around him and the 

wounded men in the rear of his car, he remarked that the 

thrill of being under fire had been pleasurable. However, 

after his thrill seeking brought him to a station closer to 

the front lines, his sense of adventure was extinguished 

by the horrors of war. After noting a French soldier died 

in the rear of his ambulance, he wrote that “...I prefer to 

forget that...Those who went through the horrors of war 

never want to talk about it.”1

Despite Service’s contention that he did not want to talk 

about war’s horrors, his writing became his venue for 

exploring his emotional reactions to combat. What is 

most striking about his autobiographical writing is the 

fact that Service developed both adaptive and idealistic endurance strategies to survive the horrors of war. 

On the one hand, he wrote that “...we became very callous, grumbling if brains or guts soiled the car. We were 

sorry for the poor devils but saw so many they were like 

shadows.”2

 In essence, he learned to treat his wounded 

charges as ‘shadows’ in order to avoid absorbing the 

emotional impact of carrying the dying on a daily basis. 

Conversely, Service also found solace in his mission. In 

his 1921 compilation of poetry and fictional prose, Ballads of a Bohemian, Service’s narrator is an ambulance 

driver like the author. The protagonist notes that “My 

only consolation is that the war must soon be over, and 

that I will have helped.”3

Thousands of ambulance drivers developed similar survival strategies as they volunteered in a war-torn Europe. 

Droves of Americans volunteered in varying capacities 

in different fronts in the years preceding their country’s 

declaration of war on Germany in 1917. Individual reasons for this massive volunteerism varied widely. Historian George Plimpton categorizes volunteer ambulance 

drivers in three groups: humanitarians, pacifists, and adventure seekers.4

 Historian Arlen J. Hansen asserts that 

many of these 3,500 documented volunteer ambulance 

drivers that enlisted before the American entry in the 

war were Ivy Leaguers who joined European armies out 

of a romantic affection for France, or even to be closer to 

family and friends in Europe. He asserts this volunteer 

fervor later spread throughout the middle class as well.5

Whatever their reasons for enlisting before 1917, American ambulanciers faced untold horrors in what was a 

Enduring for the Patients’ Sake:

 The Emotional Experiences and

 Endurance of American Ambulance

 Drivers in World War I

Benjamin Cameron Schaffer

University of New Hampshire

2016

Spurgeon Neel Annual Award Winner

The Army Medical Department Museum Foundation sponsors the Spurgeon Neel Annual 

Award competition for the original essay that best exemplifies the history, legacy, and 

tradition of the US Army Medical Department. The following essay by Benjamin Cameron 

Shaffer was selected as the best submission of the 2016 competition.
116 http://www.cs.amedd.army.mil/amedd_journal.aspx

new kind of war. In addition to the more than 8 million 

deaths in the conflict, 21 million soldiers were wounded. 

On average, about 6,000 men died every day.6

 While 

common soldiers experienced the carnage of machine 

guns and gas attacks first hand, the men evacuating 

them from the field also dealt with shellfire, the stress 

of rescuing casualties and the shocking aftermath of the 

bloodshed.

In the face of carnage, American ambulance drivers had 

disparate emotional and psychological reactions to their 

hellish surroundings. Ana Carden-Coyne, a historian of 

the British medical services, argues that morbid wounds 

“deeply affected even the most hardened surgeons who 

contemplated their patients’ grim futures...” Despite the 

horrors British medical officials faced, Carden-Coyne 

asserts that the only way they could deal with the suffering around them was “simply to get on with their jobs, 

allowing work to subsume their feelings.”7

 Despite her 

focus on British medics, Carden-Coyne’s argument for 

the persistence of British doctors and nurses easily applies to the thousands of American ambulance drivers in 

the Allied armies. The concept of forbearance through 

wartime trauma is evident in scores of published journals, poems, memoirs, and even autobiographical novels 

written by American medics both during and after the 

conflict. Ultimately, American ambulance drivers in the 

First World War not only faced extreme emotional trauma and physical dangers, but developed adaptive and 

idealistic forbearance strategies to survive their ordeal. 

Whereas ambulanciers learned to inure themselves to 

the horrors of war, they also developed positive survival 

strategies which centered on the critical value of their 

work in saving the wounded and dying.

II. Historiography of “Wartime Endurance” and

Methodology

This article is inspired by two current trends in the historiography of the First World War: the study of “troop 

endurance,” and the study of the cultural and emotional 

aspects of military medicine during the Great War. A 

recent anthology produced by World War I scholars in 

New Zealand—Endurance and the First World War: 

Experiences and Legacies in New Zealand and Australia—is a concise study on the topic of ‘endurance’ 

during the Great War.8

 In the volume, historian David 

Monger asserts that endurance’s presence has “occupied 

a quiet, yet underlying and central place in the historiography of” the war.9

 Out of the several authors Monger 

lists that have directly and indirectly covered endurance, Stephen Loveridge—author of the article “Seeing 

Trauma as Sacrifice: The Link between ‘Sentimental 

Equipment’ and Endurance in New Zealand’s War Effort,”—Michael Roper—author of The Secret Battle: 

Emotional Survival in the Great War—and Alexander 

Watson—author of Enduring the Great War: Combat, 

Morale and Collapse in the German and British Armies, 

1914-1919—have all particularly inspired this current 

study on the endurance of medics. Watson’s method of 

tracing battlefield endurance among troops amidst a sea 

of calamities will be repeated here.10

At the same time, this article is also influenced by current studies on the emotional world of Allied medics11

during World War I—particularly the work of Christine 

Hallett, author of Containing Trauma: Nursing Work in 

the First World War. While Hallett’s stated purpose is to 

broadly “uncover the hidden world of nursing work” in 

the war, her contention that nurses were forced to “contain their own emotions” lest they fail to be “of much use 

to [their] patients” is particularly relevant to this study.12 

Finally, Arlen J. Hansen’s suggestion that ambulance 

drivers’ resilience was partially based in the “intrinsic 

worthiness” of their job will be explored throughout this 

study.13 Out of all the scholarly work consulted in this 

study, Hansen comes to the closest to actually analyzing 

the endurance strategies of American ambulance drivers. All in all, this study seeks to extend groundbreaking 

arguments for World War I endurance to ambulanciers 

on the front line.

III. The Life of an Ambulancier

Ambulanciers were among the first Americans involved 

in the Great War. These idealistic young men mostly 

came from Ivy League universities, and included future 

authors of the Lost Generation such as Ernest Hemingway and John Dos Passos. These men typically served 

in one of three ambulance corps: the “Harjes Formation” 

of banker H. Herman Harjes, Richard Norton’s “AngloAmerican Volunteer Motor-Ambulance Corps”—a unit 

that later merged with Harjes’s formation under the control of the American Red Cross—and A. Piatt Andrew’s 

“American Ambulance Field Service.”14 Although there 

are incomplete records of American women who served 

as ambulance drivers, the preponderance of drivers 

were men.15

While these sections served with distinction throughout 

the conflict in France, Belgium, and even eastern regions such as Greece, the structure of most of these organizations shifted dramatically when the United States 

entered the war. The United States Army Ambulance 

Service—what Jeffrey C. Larrabee calls the “vanguard 

of the American army”—arrived in France before most 

American troops in the summer of 1917. While this organization was created by Washington to help the French 

Army, the American Expeditionary Force’s ambulance 

system was separate, and not nearly as effective as the 

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USAAS.16 Despite heavy protest from former section 

leaders—especially from Richard Norton—the three 

main ambulance groups were absorbed by the USAAS. 

One of the biggest complaints from the formerly-civilian drivers who continued on was the destruction of the 

“volunteer spirit” that came with army enlistment.17

The experiences and trials of the men in any ambulance 

section varied year by year, and from station to station. 

Wherever they served, they excelled in their primary 

task: quickly transporting the wounded to aid stations 

and evacuation centers. One unit of Andrew’s American Field Service (formerly the American Ambulance 

Field Service) was reported to have evacuated over 

56,000 wounded troops between 1915 and the war’s end 

in 1918.18 Though experiences and procedures in hospital transport obviously varied from location to location, 

Hansen maintains the American volunteers typically 

had two tasks: “jitney duty” and front line duty. Initially, 

these volunteers were permitted to take part only in “jitney” transport, which involved the transport of blessés 

(the French term for wounded soldiers) from hospital to 

hospital or from transport trains and ships to hospitals. 

As the war progressed, ambulance drivers began “working the front lines.” After a soldier was wounded, he 

was typically taken by his comrades to a first aid post, 

and from there stretcher-bearers carried him to an “advanced dressing” clinic where doctors performed basic 

operations. From here, ambulance drivers carried the 

blessés to triage stations and evacuation hospitals.19

Ambulancier James R. McConnell gives a snapshot of 

the dangerous duties these drivers had on an average 

day during the Lorraine Offensive in 1915. McConnell 

wrote: “It is difficult to take any one day’s work and describe it...for with us all days are so different...but the 

action and experiences, which add the color, are never 

alike...” He then laid out a prototypical schedule, beginning with breakfast and a discussion of German shrapnel and shell that came in the night before. Then the men 

take up their various duties. While some men are held 

in reserve, others evacuate the wounded from various 

posts both near and far. At the poste de secours (dressing station), McConnell would pick up French blessés. 

Artillery fire was intermittent at these posts, with light 

damage from shrapnel and heavy casualties from shells. 

He wrote that his section had several close calls despite 

not having lost any drivers yet.

The last of the postes was “in the very line of fire,” 

where drivers would have to watch out for craters in the 

road and frequently take cover in “bomb-proofs.” With 

the wounded in the car on the trip to the evacuation hospital, drivers started to take notice of shell damage, but 

never dwelled too long on the risks they took.20 Despite 

the intensity of these duties, Hansen contends that relatively “little of the drivers’ time was spent under fire 

at the front.” Young ambulance drivers spent upwards 

of 75% of their time ‘En Repos.’ Activities while ‘En 

Repos’ ranged from car repair to adventuring around 

Europe. Another favorite activity was writing letters or 

in journals.21 While the drivers spent most of their time 

off the battlefield, it was during these short missions that 

drivers faced many of the same anxieties and emotional 

crises as the soldiers in the trenches.

V. Emotions and Sources

In the wake of constant personal danger, the emotional 

reactions of American ambulance drivers to their environments varied by personal experience and location. 

Here it will be necessary to address a few major themes 

in emotional studies of men during the war and of emotions in wartime memoirs. Perhaps the first consideration that must be made is that the ambulanciers grew 

up in a society that valued stoicism and a “stiff upper lip” 

when undergoing trauma. Ana Carden-Coye writes that 

physical “endurance was built into military and civilian codes of masculinity—whether located in the British 

‘stiff upper-lip’ or new world images of American, Australian, and Canadian frontiersmen whose hardy bodies 

had supposedly toiled with stoic silence.” She also notes 

that among individual British doctors, “triumph over 

adversity was the main narrative of British medical and 

surgical innovation.”22 While Carden-Coyne focuses on 

physical pain, the same stoicism can be applied to men’s 

emotions as well.

Carden-Coyne and other historians have also noted the 

ability of soldiers to share their emotional responses to 

horrors in various capacities. In her study on British 

soldiers and masculinity during the war, Jessica Meyer 

argues that while British soldiers’ writings rarely “comment directly on masculinity as a specific aspect of 

identity,” their accounts reflect on how they understood 

themselves to be men, both physically and emotionally.”23 While this article does not extensively examine the 

gendered experiences of ambulanciers, it is important 

to note how reactions to combat could play into their 

perceptions of manhood—and ultimately their will to 

persevere through trauma.

Michael Roper contends that although British soldiers 

experienced a multitude of emotions, fear and dread 

were constantly on the mind of most participants. Roper 

writes that in response, these men were caught between 

unprocessed emotions and the aforementioned rule of 

the “stiff upper lip.” As we will see, the emotional responses of medics to combat were also very complex 
118 http://www.cs.amedd.army.mil/amedd_journal.aspx

and largely rooted in fear. In terms of sources, Roper 

warns that retrospective memoirs “are generally more 

reflective about the emotional experience of the war” 

than letters or journals.24 Some of the following accounts are postwar memoirs, but the raw emotional experiences recorded in the memoirs often match the more 

contemporary accounts.

One problem that does arise with wartime publications 

is the issue of subjectivity and writing for an audience. 

Arlen J. Hansen notes the compilations, With the American Ambulance Service in France25 and Friends of 

France, were both recruitment books inspired by the administration of the American Field Service. This genre 

tended to “make ambulance work sound gloriously exciting and grandly humanitarian” for recruitment and 

funding purposes.26 Nevertheless, these sources provide 

valuable insights into the emotional journeys of ambulanciers. Authors such as Jessica Meyer have examined 

the emotional world of soldiers categorically in chapters 

with titles such as “Fear,” “Discomfort,” etc. Some of 

these emotional categories will be repeated here. Broadly speaking, some of the most intense emotions ambulance drivers experienced during their service included: 

intense fear, exhaustion, and severe emotional trauma.

Fear

Fear is one of the most primal and intense emotions that 

ambulance drivers, doctors, nurses, civilians and soldiers experienced together. The closer to the front an 

individual was, the more intense their record of fear will 

be. Here it is important to distinguish between ‘fear’ and 

‘anxiety’ in battle. According to military psychologist 

and Vietnam veteran Walter F. McDermott, fear comes 

from an “external threat” (i.e. a foe). However, after the 

battle is over, anxiety—an internal form of fear—arises 

from the memories of combat. McDermott contends that 

Post Traumatic Stress Disorder, though partially psychological, often emerges from the body’s “stress response 

system continually being overstimulated,” and forms of 

‘anxiety’ often emerge as symptoms.27

Fear under fire is clearly seen in Robert Whitney Imbrie’s 1918 memoir, Behind the Wheel of a War Ambulance. Imbrie joined Andrew’s AFS at the end of 1915, 

when his section joined the French Army at Beauvais. 

Imbrie’s first experience with shell fire came in the early 

winter of 1916 in the village of Julzy. In a journal entry 

from 10 February, he wrote that “I am unable to...describe my sensations and I question whether a trained 

psychologist would be much better off. There is something disturbing about shell fire...I do not believe I was 

especially frightened; my feelings were more of curiosity.” He later wrote that “Perhaps I was not frightened 

by those first shells...but as time went on, and I saw the 

awful, destructive power of shell fire...the realization of 

their terribleness became mine.” Even after a “year and 

a half of war” he was “no more reconciled to shellfire 

than at first,” and did not think one could become inured 

to a fear that grew worse each time.28

Fear was also recorded by ambulance drivers after the 

United States entered the war in 1917. In his introduction to the diary of USAAS driver Guy Emerson Bowerman, Jr., historian Mark C. Carnes notes that Bowerman had evolving notions of fear and courage. Carnes 

contends that Bowerman wanted to appear brave before 

his comrades. On the other hand, Bowerman—like Imbrie—came to realize no one could become fully inured 

to German shells constantly flying in.29 For instance, in 

March of 1918 he reported boldly listening to his Victrola record player while “waiting either for a shell to 

hit” his house or a “call to come.” He also remarked 

that it was niceto know the Germans might be as “badly 

scared” of French shells as he was of German artillery. 

30 That Bowerman could attest to passively waiting for 

shells while also being mortified can be substantiated by 

a postwar attempt at writing a fictional autobiography, 

in which he declared that true bravery is “being afraid 

but carrying on just the same.”31 In essence, no amount 

of exposure to shells could completely eradicate fear. 

Perhaps some level of fear of shell fire was healthy in 

order to ensure self-preservation, but intense fear could 

have psychological ramifications.

Fatigue and Mental Disturbances from War’s Horrors

Aside from fear, battle fatigue was a frequent companion 

of ambulance drivers. According to Jessica Meyer, in the 

case of British soldiers’ postwar memoirs, “exhaustion 

remained a central memory of trench life for most men.” 

Not only did fatigue “[sap] men’s morale,” but even “undermined memoirists’ perceptions of themselves as men 

both physically and emotionally.”32 Sleeplessness was a 

major factor in fatigue. AFS driver Graham Carey, one 

of the first American volunteers to work as a driver at the 

Ambulance Military Hospital at Neuilly, France, complained of the lack of sleep in a letter to his mother. He 

wrote: “At four I was so sleepy that I got into a railway 

carriage that we have on a siding there and immediately 

went sound asleep. At six-thirty, however, Richardson 

and I were waked up and given two sitting cases” to 

transport.33 As one could expect, the rampant sleeplessness drivers faced led to extreme mental fatigue. While 

taking a break from the front in Paris, AFS driver Phillip 

Sidney Rice described feeling too tired and dejected to 

even regret not going out with his friends. He was particularly upset he was too poor to afford cigarettes, the only 

remedy he had to the “nerve-racking strain of war.”34

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Exposure to the horrors of war was the primary culprit 

behind this “nerve-wracking” strain. According to Michael Roper, many soldiers who were exposed to the 

war’s horrors “suffered from periods of what has been 

called ‘battle stress,’ although never becoming incapacitated to the point where they were withdrawn from 

the line.”35 More often than not, ambulance drivers also 

suffered emotional strain without being diagnosed as a 

“shell shock” case. Historian Alexander Watson notes 

that “hunger, homesickness, exhaustion and continual 

danger” were common amongst men in the trenches, 

alongside a growing sense of the “purposelessness of 

combat.” However, Watson later argues that the most 

psychologically damaging aspect of trench warfare was 

artillery fire. This danger, as we have seen, was common for ambulance drivers.36

A driver’s most obvious exposure to the horrors of war 

was through the transport of wounded men. In a poem 

by driver Emery Pottle, the narrator tries to comfort a 

wounded man in his ambulance: “Courage, mon brave! 

We’re almost there!”/ God, how the fellow groans—/ 

And you’d give your heart to ease the jolt/ Of the ambulance over the stones.” The driver laments that only God 

knew how he would “go on, through the dreadful night,” 

and notices his charge had died. He could not wake him. 

Towards the end of the poem, he laments: “it’s just another poilu [French soldier] that’s dead;/You’ve hauled 

them every day/Till your soul has ceased to wonder and 

weep/ At war’s wild, wanton play.”37 The narrator’s horror can be seen in his attempts to wake up the wounded 

blessé and admission that he wept when first exposed to 

the carnage.

Perhaps one of the most well-known results of emotional trauma during the Great War was the phenomenon of shell shock. The term, created in 1914 by British 

psychologist Charles Meyers, was used to cover a wide 

range of disturbing physical and emotional symptoms 

after combat (i.e. vomiting and vivid nightmares). By 

1916, the term “shell shock” was dually applied to two 

different sets of symptoms: physical effects from the 

shell impact and “hysteria” caused by the horrors of 

combat. The nature of the disease was so controversial 

that doctors vehemently disagreed whether physical or 

emotional trauma led to shell shock.38 It was not until 

the 1980s that doctors realized the permanent emotional 

effects combat had on soldiers and formulated the idea 

of post-traumatic stress disorder.39

Ambulance drivers were vulnerable to shell shock as 

well, Phillip Sidney Rice being a prime example. After 

one particularly exhausting mission in Verdun in 1917, 

Rice’s commander sent him to Paris for treatment for 

“battle fatigue.”40 In his memoir, Rice noted that he 

was not alone. During the battle he noticed worn out 

stretcher-bearers with their “burdens” and that his own 

face “seemed to pinch at the cheek bones” with fatigue. 

He wrote that a “couple of our men had taken sick—had 

broken down under the strain and been sent in to Paris.” 

His symptoms had been worse than theirs, however. 

Towards the end of the battle he noticed he was “playing 

out very rapidly,” was unable to relax and could not 

sleep. After his mission, he awoke in the middle of the 

night because of a “terrible nightmare” in which he 

“went through the whole experience” of battle again.41

VII. Endurance

To survive innumerable tribulations, ambulanciers were 

forced to develop mental endurance strategies. Despite 

some similarities in coping techniques, soldiers and 

ambulanciers typically developed emotional survival 

strategies particular to their jobs. For instance, Hansen 

makes the case that American ambulanciers were both 

“in and out of the war.” They experienced the mangled 

bodies of their charges and dangerous enemy fire, but 

also had the ability to drive away from the battlefield—a 

“double perspective” that alleviated some of their emotional trauma. Hansen contends that the drivers’ belief 

in the worthiness of their jobs prevented them from 

falling into despair. However, Hansen goes too far in 

asserting that drivers rarely resorted to fatalism like 

their comrades in the trenches, and that they rarely succumbed to self-doubt. As we have seen, drivers consistently doubted their ability to persevere through trials.42 

Hansen also notes that ambulance drivers found a sense 

of strength in the perceived hardiness of their French 

allies.43 Conversely, drivers such as Service developed a 

certain callousness to the war’s horrors in order to survive their jobs. In the following examination, these two 

types of endurance will be categorized as adaptive and 

idealistic endurance.

Adaptive Endurance

Alexander Watson notes how British and German soldiers were able to inure themselves to shell fire through 

constant exposure. On the other hand, to survive, soldiers had to “learn to judge risk without being overwhelmed by it” and to make judgements based on their 

chances of survival.44 Phillip Sidney Rice—the aforementioned ambulance driver who suffered shell shock—

certainly never learned to get used to shelling, but he 

was able to evaluate the benefits and consequences of 

enduring a trial as opposed to withdrawing in fear. During a bombardment in Verdun in September of 1917, 

Rice reasoned with himself that “if I do go on and am 

hit, the agony will be over within a few minutes, but if I 
120 http://www.cs.amedd.army.mil/amedd_journal.aspx

turn back, the agony will be with me the rest of my life. 

So I put on my gas mask and drove on.”45 A sense of 

shame and duty forced him to keep driving.

Such inurement strategies were not universally effective. 

Norton-Harjes driver Edward R. Coyle also initially adopted an adaptive approach to overcoming his fear of 

shell fire. Coyle wrote that “...conditions are met in a 

more or less matter-of-fact way when one is continually 

forced to accept them. Life seems a matter of fate and 

little attention is paid to bursting shells.”46 According 

to Coyle, drivers had to passively accept the harsh reality of shelling in order to carry on with their tasks. Despite this initial adaptive approach, Coyle was ultimately 

crippled by fear during one particularly heavy bombardment in Verdun.47

While Coyle realized the futility in questioning the presence of shelling in his routine, other drivers also learned 

to develop a ‘thick skin’ to overcome their fear of artillery. During a bombardment in the summer of 1915, 

AFS driver Leslie Buswell was requested to transport 

three wounded soldiers. Buswell wrote that he “had to 

grin and bear it, but it is a horrid feeling to have to go 

out into a little street where shells are falling regularly...

and run a few yards down the street to a poste de secours where a shell has just landed and another is due 

any moment.”48 Just as with the others, Buswell realized 

the inevitability of German shells, and forced himself to 

carry on out of necessity rather than desire.

Ambulanciers also had to harden themselves to the constant presence of death and disfigured corpses. Joshua G. 

B. Campbell wrote about his unit’s emotional response 

when hospital trains came in with gas victims for the 

first time. These trains came in at the same time as “a 

hail of shrapnel,” and tested the men’s emotions severely 

despite the fact that in “months of [his company’s] work 

[they] had become so accustomed to dreadful sights and 

to suffering as to be little affected by them.”49 Although 

his company became emotional when seeing gas victims, he did admit some level of adaptation to the horrors of war in Flanders fields.

Idealistic Endurance

Despite the prevalence of adaptive coping strategies, 

ambulanciers did develop positive endurance strategies. New Zealand historian Stephen Loveridge writes 

that “...conceptions “that one’s circumstances are associated with some larger or redeeming purpose, however 

that is defined, offer the human psyche some capacity to 

reframe and control apparently impossible situations.”50

On the other hand, Arlen J. Hansen notes that despite the 

“shock of the war’s horrors,” drivers did find a sense of 

adventure and education in their work.51 Drivers internalized their mission to help the wounded, and even used 

their belief in their jobs to help alleviate personal horrors. 

Of course, self-effacing service was not always healthy 

for individual drivers. Driver Leslie Buswell wrote that 

“...of course one can never allow even a shadow, much 

less a mention of one’s own problems to appear. The personal equation practically doesn’t exist here.”52

Despite having to neglect personal needs, the ambulance service did provide a form of escapism for the 

drivers. In a reflective essay that he wrote during a spell 

of pre-mission anxiety, Phillip Sidney Rice described 

seeing the horrors that the German Army wrought upon 

France, and decided he was “glad to go on living and 

loving France.”53 While one could dismiss this essay as 

mere propaganda, it is worth noting that Rice penned 

this patriotic essay while having premonitions of his 

own demise. Even in his fear, he was able to focus on 

the larger mission at hand.

Even after the United States entry into the war, American drivers continued to adopt idealistic endurance 

strategies. In June of 1918, Guy Emerson Bowerman, 

Jr. reflected on his participation in the war in the year 

since his enlistment. He wrote that although “...war is 

a terrible thing it still has its compensations for those 

who live.” Among the benefits he listed included learning “true human values...[having] more self confidence 

initiative and courage...” He admitted the “...whole 

business is unromantic,” but appreciated that there were 

“many here with me doing the same thing that it is just as 

if...this war was a perfectly natural mode of life. Thus 

are we able to adapt ourselves...”54 In essence, despite 

the horrors of war, Bowerman found purpose in his 

work and comradery with other drivers.

Perhaps the most succinct example of finding purpose in 

one’s larger mission comes from a letter Leslie Buswell 

wrote in August of 1915. He wrote that the “...horror of 

the whole war is growing on me day by day, and sometimes when I have got into my bed or am trying to get 

a few hours’ sleep on a stretcher...the horrors of blood...

haunt me, and I feel I can hardly go through another day 

of it.” In the wake of these horrors, Buswell realized “all 

that is soon forgotten when a call comes, and you see 

those bandaged soldiers waiting to be taken to a hospital. I almost love my old car...”55 In essence, the call to 

duty distracted Buswell from even the most traumatic 

memories.

VIII. Conclusion

In conclusion, American medical personnel in World 

War I experienced varying emotional responses to the 

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circumstances around them, and developed adaptive 

and idealistic endurance strategies to survive the war. 

While adaptation to trauma merely helped these medics 

to survive the horrors around them, idealistic endurance 

allowed ambulanciers to find pride in their missions that 

transcended their trials. Some might contend that accounts published during the war were mere propaganda, 

or that postwar accounts were too far removed from the 

battlefields to be reliable. While these accusations are 

reasonable, they cannot negate the existence of sundry 

emotional accounts and survival strategies found within 

these medics’ journals, letters, and memoirs.

Unfortunately, the emotional experiences of American 

ambulance drivers and physicians have been neglected 

by historians for too long. As the centennial of the conflict has arrived and every last veteran of the conflict 

has passed away, it falls upon current social and medical 

historians to not only expand the scholarship on World 

War I endurance, but to reexamine the emotional and 

social experiences of the American medics who volunteered in foreign armies and the United States military 

during the Great War.

End Notes

1. Robert W. Service, Harper of Heaven in Robert 

W. Service: Selected Poetry and Prose. Ed, Michael Gnarowski (Toronto: Dundurn Press Limited, 

2012), pp. 324-335. Google Books eBook.

2. Robert W. Service, Harper of Heaven, p. 327.

3. Robert W. Service, Ballads of a Bohemian, (New 

York: Barse & Hopkins, 1921), p. 191. Google 

Books eBook.

4. George Plimpton, “Foreword,” in Gentlemen Volunteers: The Story of American Ambulance Drivers in 

the Great War, August 1914-September 1918 (New 

York: Arcade Publishing, 1996), p. 7. Google eBook.

5. Arlen J. Hansen, Gentlemen Volunteers: The Story 

of American Ambulance Drivers I the Great War 

August 1914-September 1918 (New York: Arcade 

Publishing, 1996), pp. XV-XVI.

6. Gordon L. Heath, “Introduction,” in Canadian 

Churches and the First World War (Hamilton: McMaster Divinity College Press, 2014). Pages not 

numbered. Google Books.

7. Ana Carden-Coyne, The Politics of Wounds: Military Patients and Medical Power in the First World 

War (Oxford: Oxford University Press, 2014), pp. 

18-37.

8. David Monger, Sarah Murray, and Katie Pickles, 

“Introduction,” in Endurance and the First World 

War: Experiences and Legacies in New Zealand 

and Australia (Newcastle upon Tyne: Cambridge 

Scholars Publishing, 2014), pp. XII-XIII. Google 

Books eBook.

9. David Monger, “Endurance and First World War 

Scholarship” in Endurance and the First World 

War, pp. 1-2.

10. Alexander Watson, Enduring the Great War: Combat, Morale and Collapse in the German and British Armies, 1914-1918 (Cambridge: Cambridge 

University Press, 2008), p. 7.

11. The term medic will be used here to describe medical personnel throughout the paper. This is not to 

be confused with the ‘combat medic’ of the Second 

World War.

12. Christine Hallett, Containing Trauma: Nursing 

Work in the First World War (Manchester: Manchester University Press, 2009), pp. 1-9.

13. Arlen J. Hanse, Gentlemen Volunteers, p. 157.

14. Arlen J. Hansen, Gentlemen Volunteers, pp. 

XIV-XVII.

15. Arlen J. Hansen, Gentlemen Volunteers, p. 204.

16. Jeffrey C. Larrabee, “A Tale of Two Trucks: American Casualty Evacuation in World War I,” ICON: 

Journal of the International Committee for the History of Technology, Vol. 14 (2008), pp. 123-124.

17. Arlen J. Hansen, Gentlemen Volunteers, pp. 161-179.

18. Jeffrey C. Larrabee, “A Tale of Two Trucks,” p. 128.

19. Arlen J. Hansen, Gentlemen Volunteers, pp. 

XV-XVI.

20. James R. McConnell, “The Section in Lorraine,” 

in Friends of France: The Field Service of the 

American Ambulance Described by its Members, 

ed. (Boston: Houghton Mifflin Company, 1916), pp. 

71-81.

21. Hansen, Gentlemen Volunteers, Google eBook, pp. 

140-157.

22. Ana Carden-Coyne, The Politics of Wounds, pp. 

162-183.

23. Jessica Meyer, Men of War: Masculinity and the 

First World War in Britain (Houndmills: Palgrave 

Macmillan, 2011), p. 9.

24. Michael Roper, The Secret Battle: Emotional Survival in the Great War (Manchester: Manchester 

University Press, 2009), pp. 14-20.

25. Also published as Ambulance No. 10: Personal Letters from the Front.

26. Arlen J. Hansen, Gentlemen Volunteers, pp. 53-54.

27. Walter F. McDermott, Understanding Combat Related Post Traumatic Stress Disorder (Jefferson: 

McFarland & Company, Inc., 2012), pp. 48-49.. 

Kindle eBook.

28. Robert Whitney Imbrie, Behind the Wheel of a War 

Ambulance (New York: Robert M. McBride & Co., 

1918), pp. 4-40.
122 http://www.cs.amedd.army.mil/amedd_journal.aspx

29. Mark C. Carnes, “Introduction,” in Guy Emerson 

Bowerman, Jr., The Compensations of War: The 

Diary of an American Ambulance Driver During 

the Great War (Austin: University of Texas Press, 

1983), pp. XIV-XVII.

30. Guy Emerson Bowerman, Jr., The Compensations 

of War, pp. 67-68.

31. Mark C. Carnes, “Introduction,” in Guy Emerson 

Bowerman, Jr. The Compensations of War, p. XV.

32. Jessica Meyer, Men of War, pp. 131-132.

33. Graham Carey to His Mother, 10 January 1915, in 

Lyn Macdonald, ed. The Roses of No Man’s Land

(London: Michael Joseph Ltd, 1980), pp. 63-65.

34. Rice, An American Crusader, p. 81.

35. Michael Roper, The Secret Battle, p. 247.

36. Alexander Watson, Enduring the Great War, pp. 

20-27.

37. Emery Pottle, “Un Blesse A Montauville” in 

Friends of France, pp. 136-138.

38. Christine Hallet, Veiled Warriors: Allied Nurses of 

the First World War (Oxford: Oxford University 

Press, 2014), pp. 95-98. Kindle eBook.

39. Walter F. McDermott, “History of Combat Disorders,” in Understanding Combat Related Post Tramautic Stress Disorder, Kindle eBook version.

40. Arlen J. Hansen, Gentlemen Volunteers, p. 69.

41. Phillip Sidney Rice, American Crusader at Verdun, 

pp. 72-79.

42. Arlen J. Hansen, Gentlemen Volunteers, pp. 155-157.

43. Arlen J. Hansen, Gentlemen Volunteers, pp.

44. Watson, Enduring the Great War, Location 2647, 

Kindle eBook Version.

45. Phillip Sidney Rice, American Crusader, pp. 76-78.

46. Edward R. Coyle, Ambulancing on the French 

Front (New York: Britton Publishing Company, 

1918), pp. 41-42.

47. Edward R. Coyle, Ambulancing on the French 

Front, pp. 81-83.

48. Buswell, Ambulance No. 10: Personal Letters from 

the Front (Boston: Houghton Mifflin Company, 

1916) pp. 99-101. Google Books eBook.

49. Joshua G.B. Campbell, “The Section in Flanders,” 

in Friends of France, pp. 121-122.

50. Steven Loveridge, “The Link Between Sentimental 

Equipment and Endurance in New Zealand’s War 

Effort,” in Endurance and the First World War: 

Experiences and Legacies in New Zealand and 

Australia, eds. David Monger, Sarah Murray, and 

Katie Pickles (Newcastle upon Tyne: Cambridge 

Scholars Publishing, 2014), p. 54.Google Books 

eBook.

51. Arlen J. Hansen, pp. 197-202.

52. Leslie Buswell, Ambulance No. 10, p. 87.

53. Phillip Sidney Rice, American Crusader, pp. 

98-101.

54. Guy Emerson Bowerman, Jr. The Compensations 

of War, pp. 107-108.

55. Leslie Buswell, Ambulance No. 10, p. 98.

Bibliography

Primary Sources

Bowerman, Guy Emerson, Jr., The Compensations of 

War: The Diary of an American Ambulance Driver During the Great War. ed, Mark C. Carnes. Austin: University of Texas Press, 1983.

Buswell, Leslie. With the American Ambulance Field 

Service in France. Houghton Mifflin: 1915. Google 

Books eBook.

Ambulance No. 10: Personal Letters from the Front. 

Boston: Houghton Mifflin Company, 1916. Google 

Books eBook. Accessed 27 December 2015.

Coyle, Edward R. Ambulancing on the French Front. 

New York: Britton Publishing Company, 1918. Accessed 

November 26, 2015. Archive.org. https://archive.org/

stream/ambulancingonfre01coyl#page/n7/mode/2up.

Howe, M.A. DeWolfe, ed. The Harvard Volunteers in 

Europe: Personal Records of Experience in Military, 

Ambulance, and Hospital Service. Cambridge: Harvard 

University Press, 1916.

Richard Norton to Eliot Norton, 7 June 1915.

Imbrie, Robert Whitney. Behind the Wheel of a War Ambulance. New York: Robert M. McBride & Co., 1918. 

Accessed November 21, 2015 Archive.org. https://archive.org/stream/behindwheelofwar00imbr#page/n7/

mode/2up.

Macdonald, Lyn ed. The Roses of No Man’s Land. London: Michael Joseph Ltd, 1980.

Graham Carey to His Mother, 10 January 1915.

Piatt, Andrew A, ed. Friends of France: The Field Service of the American Ambulance Described by its Members. Boston: Houghton Mifflin Company, 1916.

McConnell, James R. “The Section in Lorraine”.

Emery Pottle, “Un Blesse A Montauville”.

Sidney Rice, Phillip. An American Crusader at Verdun

(Princeton: Princeton University Press, 1918. Google 

Books eBook.

ENDURING FOR THE PATIENTS’ SAKE: THE EMOTIONAL EXPERIENCES AND ENDURANCE

OF AMERICAN AMBULANCE DRIVERS IN WORLD WAR I
January – June 2017 123

THE ARMY MEDICAL DEPARTMENT JOURNAL

Service, Robert W. Harper of Heaven in Robert W. Service: Selected Poetry and Prose, ed. Michael.

Gnarowski. Toronto: Dundurn Press Limited, 2012. 

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Ballads of a Bohemian. New York: Barse & Hopkins, 

1921. Google Books eBook.

Rhymes of a Red Cross Man. Barse & Hopkins: 1916. 

Google Books eBook.

Secondary Monographs

Carden-Coyne, Ana. The Politics of Wounds: Military 

Patients and Medical Power in the First World War. Oxford: Oxford University Press, 2014.

Hallett, Christine. Containing Trauma: Nursing Work in 

the First World War. Manchester: Manchester University Press, 2009.

Veiled Warriors: Allied Nurses of the First World War. 

Oxford: Oxford University Press, 2014. Kindle eBook.

Hansen, Arlen J. Gentlemen Volunteers: The Story of 

American Ambulance Drivers I the Great War August 

1914-September 1918. New York: Arcade Publishing, 

1996.

Heath, Gordon L. Heath, Canadian Churches and the 

First World War. Hamilton: McMaster Divinity College 

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McDermott, Walter F. Understanding Combat Related 

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Meyer, Jessica, Men of War: Masculinity and the First 

World War in Britain. Houndmills: Palgrave Macmillan, 

2011.

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in Endurance and the First World War: Experiences 

and Legacies in New Zealand and Australia (Newcastle 

upon Tyne: Cambridge Scholars Publishing, 2014), pp. 

XII-XIII. Google Books eBook.

Loveridge, Steven “The Link Between ‘Sentimental Equipment’ and Endurance in New Zealand’s War 

Effort”.

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in the Great War. Manchester: Manchester University 

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Scholarly Article

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(2008).
124 http://www.cs.amedd.army.mil/amedd_journal.aspx

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