Skip to main content
NCBI home page
NIHPA Author Manuscripts logoLink to NIHPA Author Manuscripts
. Author manuscript; available in PMC: 2017 Nov 1.
Published in final edited form as: J Forensic Sci. 2016 Oct 5;61(6):1578–1587. doi: 10.1111/1556-4029.13189

Fluorescence Imaging of Posterior Spiracles from Second and Third Instars of Forensically-important Chrysomya rufifacies (Diptera: Calliphoridae)*

Danielle Flores 1,3, Amy L Miller 3, Angelique Showman 1,2, Caitlyn Tobita 1, Lori MN Shimoda 1, Carl Sung 1, Alexander J Stokes 4, Jeffrey K Tomberlin 5, David O Carter 3, Helen Turner 1
PMCID: PMC5118101  NIHMSID: NIHMS813463  PMID: 27706817

Abstract

Entomological protocols for aging blow fly (Diptera: Calliphoridae) larvae to estimate the time of colonization (TOC) are commonly used to assist in death investigations. While the methodologies for analysing fly larvae differ, most rely on light microscopy, genetic analysis or, more rarely, electron microscopy. This pilot study sought to improve resolution of larval stage in the forensically-important blow fly Chrysomya rufifacies using high-content fluorescence microscopy and biochemical measures of developmental marker proteins. We established fixation and mounting protocols, defined a set of measurable morphometric criteria and captured developmental transitions of 2nd instar to 3rd instar using both fluorescence microscopy and anti-ecdysone receptor Western blot analysis. The data show that these instars can be distinguished on the basis of robust, non-bleaching, autofluorescence of larval posterior spiracles. High content imaging techniques using confocal microscopy, combined with morphometric and biochemical techniques, may therefore aid forensic entomologists in estimating TOC.

Keywords: forensic science, forensic entomology, post-mortem interval, calliphoridae, fluorescence microscopy

Utilizing insects associated with human remains to determine forensically relevant information is a well-accepted practice (1–7). Insects can serve as valuable physical evidence for several different types of investigation. Insects can be used to associate people and objects with a particular habitat (8), serve as a source of suspect DNA in sexual assault investigations (9) and indicate the presence of drugs in decomposing remains (3). Another common use of insects is to estimate the time of colonization, which could infer the postmortem interval, the amount of time that has lapsed since death (8), given select assumptions (10). This is often achieved by establishing the age of blow fly (Diptera: Calliphoridae) larvae, which is significantly correlated to temperature. This process allows a forensic entomologist to estimate a minimum time of colonization (TOC), which serves as an estimate of minimum postmortem interval (10).

Blow flies (Diptera: Calliphoridae) are primary colonizers of vertebrate remains, including humans (11–13). In most cases, blow fly immature development includes an egg, three instars and the pupa (11–15). Key morphological features used to identify the species and instar are located in the spiracular openings found on the posterior end of the larva (16–18). Identification of growth stages can be accomplished by counting the number of spiracular slits present in each spiracle (18). As time progresses the number of these spiracular slits increases so that one slit per spiracle indicates 1st instar, two slits per spiracle indicates 2nd instar, and three slits per spiracle indicates 3rd instar. The spiracle size of a blow fly is relatively small to the naked eye. Light microscopy resolution limits observation of morphological characteristics, in turn limiting the resolution of TOC estimates. Current methods to distinguish between larval stages are viewing the spiracles under low-resolution light microscopy (18). Viewing the larvae using SEM provides far greater resolution but is technically challenging and resource-intensive (19).

In this study we postulated that confocal or fluorescence microscopic analyses of blow fly larvae (19–21) could improve estimates of TOC through: providing new parameters for distinction of larval stage, such as the intensity and distribution of auto-fluorescent proteins or secondary metabolites, and increasing spatial resolution of morphological features. We conducted a pilot study using epifluorescence and confocal analysis of Chrysomya rufifacies (Macquart) (Diptera: Calliphoridae) larvae, which frequently are encountered on human remains in select regions of the world (11, 12) reared under laboratory conditions. We established fixation and mounting protocols, defined a set of measurable morphometric criteria and captured developmental transitions of 2nd instar to 3rd instars. These data show that instar stages can be distinguished on the basis of epifluorescence microscopy and confocal microscopy. We propose, and demonstrate, confirmatory biochemical analyses as a complimentary approach. Fluorescence microscopy may therefore provide an important method in the suite of approaches available for forensic entomologist to assist in estimating TOC.

Materials and Method

Larvae Rearing and Mounting Preparations

Raw chicken (~100 g) was placed in an outdoor setting on the campus of Chaminade University of Honolulu, Oahu, Hawaii (21.3° N, 157.8° W) for 1 h per day for three days. Mean daily temperature was 30 °C. Flies positively identified as C. rufifacies that were attracted to this sample were captured and used to initiate a colony. The colony was maintained on beef liver under controlled conditions in a 60 cm × 60 cm × 60 cm rearing chamber with 1.9 cm vermiculite bed maintained at an average daily temperature of 20.4 °C (HOBO Pro V2 U23-001 Optic datalogger, Bourne, MA). Developing larvae were removed at intervals, heated at 80 °C for 5 min and fixed in 70% ethanol (17, 22–26). Fixed larvae were dissected under a standard dissecting microscope (Nikon, Melville, NY) initially with a transversal incision across the 11th abdominal segment. Pinning and mounting was performed under an ethanol drop to avoid dehydration, and further dissection removed any intestinal or other structural material. Mounting under a glass coverslip used 100 μl of partially dried Crystal Mount (Electron Microscopy Sciences, Hatfield, PA) to avoid sample compression.

Chemicals and Reagents

All standard chemicals used in the study were from Sigma Chemical Company (St. Louis, MO) or VWR (Radnor, PA) unless otherwise specified.

Imaging

Images were taken with a Nikon Ti-U inverted scope. A Nikon (Nikon, Melville, NY) D-Eclipse C1 system was used for laser scanning confocal microscopy with Nikon EZC1 software. Available laser lines in FITC (fluorescein isothiocyanate), TxRed (Texas Red) and Cy5 (Cyanine 5) channels were supplied by a 488 nm 10 mW solid state laser, a 561 nm 10 mW diode pump solid state (DPSS) laser and a 638 nm 10 mW modulated diode laser. A Nikon DS-Qi2 monochrome camera was used for taking epifluorescence images using 4 filter sets: DAPI (Alexa 350), FITC (Alexa 488), TxRED (Alexa 568), Cy5 (Alexa 647), with all Alexa conjugates supplied by Molecular Probes (Eugene, OR). Nikon NIS-Elements Advanced Research software was used for acquiring the epifluorescence images and the analysis of both the confocal and epifluorescent images. Pinhole size for all images was 60 μm. Epifluorescent and bright field images were taken with a DS-Qi2 camera, with resolution at 1636×1088 at 14 bit. Images were calibrated using a stage micrometer in conjunction with the 20x and 40x objectives used. Images were analysed in NIS Elements.

Spiracle Morphometric Measurements

Confocal 2nd and 3rd instar spiracle images were selected and measured using Annotations and Measurements tools in NIS Elements-AR Software (Nikon, Melville, NY). Multiple Z disc of each 2nd and 3rd instar image were selected and used to manually measure length, area and number of the inner spiracular processes of that spiracle in each channel (FITC-515 nm, TxRed-590 nm and Cy5-650 nm).

Protein Extraction and Western Blots

Unfixed larvae (~0.1g) were lysed (ice/30 min) in 500μl of lysis buffer (1% (w/v) IGEPAL, 50mM Hepes pH 7.4, 250mM NaCl, 40mM NaF, 10mM iodoacetamide, 0.5% (w/v) Triton X100, 1mM phenylmethylsulfonylfluoride, 12.5mM sodium pyrophosphate 500 μg/ml aprotinin, 1.0 mg/ml leupeptin and 2.0 mg/ml chymostatin). Lysates were clarified (17,000g, 20 min) and acetone precipitated (1.4 volumes acetone, 1h/–20°C, followed by 10,000g/5 min). Protein determination used a Dc protein determination kit (BioRad, Temecula, CA) with Bovine Serum Albumin as a standard. Protein was resolved by 10% reducing SDS-PAGE in a modified Laemmli buffer and electro-transferred to PVDF in 192mM glycine, 25mM Tris (pH 8.8). Membranes were blocked (5% non-fat milk, 5% BSA and fish skin gelatin (Sigma, St. Louis, MO) in PBS, 1h, 22–25°C) and probed (primary antibodies in PBS/0.05% Tween-20/0.05% NaN3, 16 h/4°C). Primary antibodies tested were anti-ultraspiracle (Abcam, Cambridge, MA), anti-ecdysone receptor (Abcam, Cambridge, MA), anti-cofilin (Abcam, Cambridge, MA), anti-β-tubulin (Cell Signaling Technology, Danvers, MA), and anti-β-actin (Abcam, Cambridge, MA). Developing antibodies comprised anti-rabbit or anti-sheep IgGs conjugated to horseradish peroxidase (Amersham) at 0.1 micrograms/ml in PBS/0.05% Tween-20 (45 min/RT). Signal was visualized using ECL (Amersham) and Kodak BioMax film. Films were scanned at >600 dpi and quantification was performed using ImageJ (NIH, Bethesda, MD).

Statistical Analysis

Results are shown as mean ± SD. Data met parametric criteria. Therefore, statistical significance was determined based on Student’s t test or analysis of variance (ANOVA; GraphPad Prism 6 v6.02; La Jolla, CA). Adjacent to data points in the respective panels, significant differences were recorded as *p < 0.05, **p < 0.01, or ***p < 0.001, where n = 3. #= not calculated.

Results

Significant, Bleaching-resistant Intrinsic Fluorescence Enables High-resolution Confocal Imaging of Chrysomya Rufifacies Posterior Spiracles

Figure 1A (left panel) shows an example larva, with the posterior spiracles clearly visible. Relative sizing of first, second and third instar larvae are shown in Figure 1A, center panel. Figure 1A (right panel) presents visible light imaging of major features of the posterior spiracle. After sectioning and mounting, we first asked if the posterior spiracle sections emitted robust autofluorescence (27) sufficient for lengthy (multiple minute) serial sectioning using confocal laser scanning microscopy. Strong fluorescence was noted in the FITC (λex 488 nm, λem 515/30 nm, Figure 1B plate ii), TxRed (λex 561 nm, λem 590/50 nm, Figure 1B plate iii) and Cy5 (λex 638 nm, λem 670 nm, Figure 1B plate iv) channels. Less intense signal was noted in the DAPI channel (λex 358 nm, λem 461 nm). Bleaching times were compatible with prolonged imaging. For example, we noted <10% loss in fluorescence over a 16 min illumination period with a medium pinhole at 40% laser strength in the FITC channel. We also noted (Figure 1C) that fluorescence intensity (assessed using intensity surface plotting) in the three channels was differentially distributed between z discs, highlighting different structures. We performed serial sectioning and 3D reconstruction, viewing these reconstructions from both medial and caudal sides of the posterior spiracle (orientation shown schematically in Figure 1D). Figure 1E shows a 3D reconstruction of serial images through the posterior spiracle of a third instar larva in three fluorescence channels, and merged. Figures 1E–F present medial and basal imaging projections of z series, reassembled into maximum intensity projections and rendered in 3D. Figure 1G exemplifies the level of resolution possible by virtue of the robust fluorescence that enables long imaging series without loss of signal. Here, a structure located between spiracular slits (the spiracular plate (28) was targeted for focused imaging at high magnification (100X) in a detailed z series (120 serial sections at 150 nm step size). This structure is located in Figure 1G (iv) below. The resulting images clearly offer high resolution and considerable potential diagnostic and scientific interest.

FIG. 1. Confocal fluorescence imaging of C. rufifacies larvae.

FIG. 1

FIG. 1

FIG. 1

FIG. 1

FIG. 1

Open in a new tab

A. Conventional imaging of instars and posterior spiracles. Left Panel. Photograph of 3rd instar larva from C. rufifacies, showing posterior spiracles. Scale bar is 0.5 cm. Center panel. Size comparison of first, second and third instar larvae. Scale bar is 1.0 cm. Right panel. Light microscope image (40x) of 3rd instar larva from C. rufifacies, showing posterior spiracles.

B. Confocal imaging of intrinsic fluorescence of C. rufifacies posterior spiracle at third instar. (i) 20x image, confocal z series of 53 sections of 200nm vertical depth, merged and 3D reconstructed using maximum intensity projection from sequential images taken in the FITC, TXRed and Cy5 laser lines. Merged images are pseudocolored green (FITC channel), orange red (TxRed channel) and Deep Red (Cy5 channel), with co-incident fluorescence appearing as yellow. (ii–iv) Single z discs (step 11/53) from the series in (i), presented pseudocolored green (FITC channel), orange red (TxRed channel) and deep red (Cy5 channel). Scale bar is 200 microns.

C. Intensity surface plots of differential fluorescence intensity and spatial distribution throughout posterior spiracle z series and between fluorescence channels. Z discs of the step indicated at left (8–40) of a 53 image z series were analysed for the spatial distribution of fluorescence intensity (intensity surface plot, ISP) and visualized in three fluorescence channels using matched y-axis scaling. Merged images of the relevant z disc are shown at left for reference. Merged images are pseudocolored green (FITC channel), orange red (TxRed channel) and Deep Red (Cy5 channel), with co-incident fluorescence appearing as yellow. Scale bar is 200 microns.

D. Schematic of image analysis for Figure 1E.

E, F. Basal and medial views of 3D reconstructions of serial third instar confocal images in three fluorescence channels. 20x image, confocal z series of 53 sections of 200nm vertical depth, merged and 3D reconstructed using maximum intensity projection from sequential images taken in the FITC, TXRed and Cy5 laser lines, viewed as merged three channel images in basal and medial projections (E, i and ii), and as separated FITC (E, iii and iv), TxRed (F, i and ii), and Cy5 (F, iii and iv) channel images in basal and medial projections. Merged images are pseudocolored green (FITC channel), orange red (TxRed channel) and Deep Red (Cy5 channel), with co-incident fluorescence appearing as yellow. Scale bar is 200 microns.

G. High-resolution imaging of interspiracular slit structure facilitated by robust, non-bleaching intrinsic fluorescence.

Three serial z discs at positions 56 (i), 57 (ii) and 58 (iii) of a 120 image series (step size 150 nm) were captured using a 100X objective in the FITC fluorescence channel. Images were rendered using maximum intensity projection and a reference image (iv) to locate the highlighted structure is shown. Scale bar is 10 microns.

Morphometric Schema for Comparative Analysis of Chrysomya Rufifacies Posterior Spiracle Images

Throughout these imaging experiments, we noted specific structural features including the ecdysial scar, rima, peritreme area, spiracular hairs and intraspiracular processes (19, 25, 26) that were differentially apparent between fluorescence channels, suggesting that imaging protocols can be targeted to highlight desired structural features in a straightforward manner. The level of resolution and detail in the resulting images supports the premise that confocal imaging could be used to discern larval developmental stage at a fine level of resolution. However, in order for fluorescence analysis to become standardized and useful for developmental staging, we reasoned that morphometric criteria that can be robustly compared, between samples and across time, needs to be specified. Quantifiable measurements that can be assessed in a reference population and demonstrated to reliably distinguish development stages will be needed if this approach is to have utility in the forensic community. As a first step towards this goal, we developed criteria for posterior spiracle morphometry defined by basic parameters (e.g., length, width) and feature-specific parameters. Figure 2A (i) locates the morphometry parameters used in this study on an example confocal z disc of a 3rd instar posterior spiracle.

FIG. 2. Confocal fluorescence imaging of C. rufifacies larvae.

FIG. 2

Open in a new tab

A. Morphometry of third instar imaging. (i) An example third instar z disc is shown, with morphometric parameters (defined in Table I) superimposed. (ii–vii) Magnified views of spiracular slit (ii, iii), intraspiracular processes (iii), spiracular hairs (iv, v) and ecdysial scar (vi, vii) provided to orient the morphometry displayed in Table I.

B. Tukey analysis. Plot of second and third instar distributions (median, 25th and 75th percentile, min and max) of measurements for parameter a (circumference, spiracular disc) across three independent samples.

We applied these morphometric criteria to the discrimination of second and third instar posterior spiracles. Table I shows second and third instar morphometry data, averaged across three independent samples. Statistically significant differences between instar measurements are indicated as follows: *p < 0.05, **p < 0.01, or ***p < 0.001. Several of the parameters measured attained statistical significance between instars, an expected result. Of interest are the ranges within instar categories. Tukey plots of one major parameter (parameter a, length of longest axis) are shown in Figure 2B to illustrate ranges of measurements between instars. Since the exact timing of oviposition and initiation of the developmental process was not matched between larvae, these measurements probably reflect a continuum of developmental changes that could be used to more precisely position larval age and hence PMI.

TABLE 1.

Morphometric analysis of 3 independent replicates of second and third instar posterior spiracular discs of C. rufifacies

Parameter 2nd Instar 3rd Instar
Spiracular disc
Circumference x 7,850 31,400***
Length, longest bisecting axis a 103 206***
Length, shortest bisecting axis b 84 196#
Area, total
Area, internal		 c
d (ratio, e) 		6,680 30,668***
Area, Peritreme f N.D. N.D.
Peritreme open, closed open closed
Spiracular Slits
Number of slits g 2 3***
Length, mean of longest bisecting axis h 66 134**
Width, mean of longest bisecting axis h′ 25 42*
Area i (mean, j) 1655 4108**
Ecdysial Scar
Area, ecdysial scar l 248 1,729**
Intraspiracular processes
Count, tips m 11 17*
Spiracular hairs
Count, origins n 6 6#
Open in a new tab

Confocal Imaging and Supporting Biochemical Approaches May Distinguish Developmental Stages in Chrysomya Rufifacies Larvae

We compared instars using imaging techniques and obtained comparison images. Figure 3A illustrates example data from a first instar, where immature spiracular slits were observed with limited visible internal structures. This limits the degree to which anything other than basic morphometry (e.g., length, width, area) of first instar slits would be useful diagnostically. Differential imaging of second and third instar posterior spiracles was straightforward, and we were able to also capture transitional events of the kind shown in Figure 3B and 3C. Here, posterior z discs of 113 serial sections at 150nm step distance were obtained (see schema in Figure 1D). Tiled z discs (Figure 3B) allow easy discrimination of mature third instar and the residual second instar spiracles, which are being absorbed into the peritreme/scar region, whereas the ecdysial scar appears non- or under-developed in the first instar. Figure 3C presents basal and medial 3D reconstructions of this z series, as maximum intensity projection with rotation.

FIG. 3. Comparative imaging of C. rufifacies instars and transition states.

FIG. 3

FIG. 3

FIG. 3

Open in a new tab

A. Summary of first instar imaging. (i) Bright field 40x image, scale bar 200 microns, of a first instar C. rufifacies posterior section. (ii) Merged fluorescent image of FITC, TXRed and Cy5 channel images of a single z disc (11 of a 22 step series, 150 nm per step) that approximately bisected the developing spiracular slits. Scale bar is 100 microns. (iii) TxRed image of z disc as in (ii), with some loose internal structure visible. Scale bar is 50 microns. (iv) A merged three-channel image of first instar spiracle z disc as in (i). Merged images are pseudocolored green (FITC channel), orange red (TxRed channel) and Deep Red (Cy5 channel), with co-incident fluorescence appearing as yellow. Scale bar 50 microns.

B, C. Second to third instar transition imaging. B. Tiled presentation of merged three-channel (FITC, TxRed, Cy5) images of 6 z discs (steps 15, 25, 33, 44, 55, 66, (i–vi, respectively), of a 113 step z series with 150 nm step size through the posterior spiracle of a transitioning second-third instar C. rufifacies larva. C. Merged three channel images of the 113 z discs reconstructed in 3D and rendered using maximum intensity projection. Basal (i) and medial (ii) projections oriented as in Figure 1D are shown. Scale bars are 50 microns.

We sought biochemical methods to provide a supporting methodology to imaging for larval age determination. We screened a number of commercially available antibodies raised against Drosophila antigens, reasoning that these might be likely to display cross-reactivity to Chrysomya proteins. Drosophila (Diptera: Drosophilidae) is the only dipteran species for which antibodies are routinely commercially available. Antigens were selected based on their likelihood of varying across developmental stages. Of the 2 antigens studied (Ultraspiracle (29) and Ecdysone Receptor (30–32)), only the Ecdysone Receptor Western blotted positively across species and varied in apparent expression levels between instars (Figure 4A). We also explored, bioinformatically, the available protein sequences that could guide selection of antibodies or the raising of new antibodies in the future. Of the 300 protein sequences currently deposited in NCBI, 273 are mitochondrial protein subunits (Cytochrome Oxidase, Cytochrome b, NADH Dehydrogenase, F1-F0-ATPase). These are highly conserved between Drosophila and Chrysomya (data not shown) but are not a priori particularly likely to be expressed specifically at certain stages of development. More than 20 other proteins listed in Table II were examined for homology to Drosophila, published or experimental evidence of variation across developmental stages, and antigenicity (Table II). Of these, commercially-available antibodies are limited to Odorant Co-receptor (Orco), Sex-lethal (Sxl) Bucoid (Bcd) and EF1-alpha, but not carbamyl phosphate synthase, setting the stage for a wider study of the biochemical utility of these markers in determining larval stage.

FIG. 4. Biochemical analysis of instar progression.

FIG. 4

Open in a new tab

A. Western blot analysis of first, second, and third instar Ecdysone Receptor expression. Post-nuclear lysates from larval samples were resolved by 10% SDS-PAGE on a large scale Hoeffer gel system, allowing the loading of 500 micrograms total protein per lane. After electrophoresis, Western blotting with 0.1 μg/ml anti Ecdysone Receptor (EcR) was performed. Predicted molecular weight of EcR (based on Drosophila protein) is 80 kDa. MW shown in kDa at left of gel.

TABLE 2.

Potential biochemical markers for instar progression based on meta-analysis of C. rufifacies protein and EST sequence representation in NCBI database

Protein Name (GI accession) Developmentally regulated? D. Melanogaster
% identity, % positive, GI accession
Odorant co-receptor
GI:384503150 		N.D. 88,93, GI:24644231
Sex-lethal
GI: 25291016; 6226775		 GDS3835
Early pupal>adult=late
pupal>larval		 65, 73, GI:281359985
Bicoid
GI: 695320128; 695320125; 695320123; 695320120; 695320118; 695320115; 695320112; 695320110; 695320107; 695320102; 695320100; 695320097; 695320051; 695320049; 695320047		 Yes 69, 75, GI:18652210
Elongation Factor 1-alpha
GI: 377648422; 397529974; 397529972; 397529970; 258537035		 GDS3835
adult=late pupal=early
pupal≫larval		 97, 98, GI:17137572
Carbamyl phosphate synthase
GI: 257050977; 258536849		 GDS3835
late pupal=early
pupal=larval≫adult		 89, 93, GI:4337094
Open in a new tab

EST, expressed sequence tag; NCBI, National Center for Biotechnology Information. GI, gene identifier; GDS, GEO dataset. N.D., not determined

Discussion

We determined that epifluorescence and confocal analyses can be used to view morphological features associated with different instars of C. rufifacies. The advanced imaging techniques applied in this study offer improved discriminatory power over the currently available level of definition available from low-resolution light microscopy. Improved resolution enables the discrimination of structural and morphological differences that change between instars and offer the prospect of staging a larvum within the intra-instar developmental time frame. In addition to these determinations of structural change during development, structure-dependent and -independent changes in fluorescence intensity profile may also be of diagnostic use. Our findings add to the discriminatory power of morphometric measurements where the following attributes of the spiracle appear to be sufficiently variant to distinguish instars: (a) size, (b) number of slits, (c) dimensions of slits, (d) slit internal structural complexity, (e) number of spiracular hairs, (f) peritreme and ecdysial scar dimensions and (g) dimensions and location of the spiracular plate. Our study reinforces and extends the work of Samerjai in tropical flesh flies, where the distance from the boundary of the spiracular compartment to the tip of the spiracular plate structure was suggested as a discriminator of larval stage (28, 33). In addition, the fluorescence intensity profiles of the spiracular structures vary spatially, temporally and between fluorescence channels, providing an untapped set of potential discriminators between stages. Clearly, the combination of morphometry, fluorescence intensity measurements and rigorous reference data sets offer the potential to improve accuracy of staging.

One key development in the field of estimating TOC will be the capacity to resolve time periods in the intra-instar periods. This is a complex undertaking that offers significant potential to improve definition of m-PMI. Our data suggest that within the enhanced resolution gained by confocal imaging, discernment of more discrete developmental stages than the instar boundaries may be possible, most notably what may be the first visualization of the inter-instar transitional period of a second to third instar larvae. Again, reference data sets become key as we move towards this goal, and in ongoing experiments we are seeking to compile the first of these reference sets with controlled rearing of larvae from C. ruffifacies, harvested hourly in sufficient numbers to offer statistical power to the experiment, and imaged according to the protocols defined here.

At a minimum, we submit that our current study documents the potential utility of capturing the inter-instar transitional period. By chance, we also happened to capture a transitioning larva at the second to third instar boundary. This serendipitous sample offered two new insights: First, that instar transitions are readily apparent microscopically, and likely represent an opportunity to state a time since colonization that mark the transitional time period at increased time resolution. Second, the apparent resolution of the second instar spiracle into the peritreme, and the brief coexistence of the two- and three-slit spiracular structures suggests a similar, visible event must take place between the first and second instar boundary. This is a time period that occurs very rapidly in larval development that instead may be able to provide an even closer estimate to PMI in fractions of an hour. Furthermore, first instar larvae appeared to lack an ecdysial scar, thus the first to second inter-instar transition may result in the formation of this morphological feature. Indeed, our initial findings are of entomological interest and deserving of more study.

Correct identification of insect species collected as evidence during forensic investigations is critical for precise and accurate determination of information important to the investigation. It is therefore important to consider potential confounding factors and caveats to the proposition that fluorescence microscopy can be useful in staging TOC (7, 12, 13). The first of these challenges will be whether genetically- or environmentally-driven variability in larvae will simply overwhelm the measurable differences between samples. For example, in blow flies, development can be highly variable depending on the species being examined and the environmental conditions in which it originated. C. rufifacies needs approximately six days to complete larval development at 28° (12), while the closely related species, Chrysomya megacephala (Fabricius) (Diptera: Calliphoridae) needs 5.4 days at 26°C (34). Regardless, these two species frequently occur in the same regions of the world and appear similar during the early portion of larval development. Failure to identify specimens correctly could result in incorrect estimates of the time of colonization and thus time of death estimates. This may mean that PCR-based determination of phylogeny based on 16S sequencing may need to accompany the staging possible through imaging protocols. Variance in development based on the tissue substrate may also be a factor. The variance in the system will be captured in future studies that establish reference sets, but if a reference set is actually required for every different locale, ecozone, or species this may be a tremendous practical barrier. Nevertheless, even if only the most variant spiracle characteristics emerge as useful because they are common between locales and across species, it is likely that the improvements in resolution (c.f., Figure 1A and B) offered by fluorescence microscopy will be useful. More broadly, applications of this type of approach may extend beyond TOC estimation, to support geoforensics and associate persons and objects with particular locations for criminal justice and defense purposes.

A second major challenge to implementation lies in cost and expertise associated with the use of fluorescence imaging. These challenges might require that local academic partners be enlisted for crime laboratories seeking to incorporate these methodologies. Compliance with International Standards Organization (ISO) and American Society of Crime Laboratory Directors (ASCLD) applicable standards would be needed, possibly necessitating the development of new standards for forensic biological imaging. Without these developments, acceptance at the evidentiary level would be highly challenging.

The robust intrinsic fluorescence of insect samples offers significant scientific opportunity within both forensic and non-forensic applications of entomological fluorescence imaging. First, it may be possible to supplement or even replace imaging using fluorescence with fluorescence spectroscopy of homogenized larval samples, which has potential application to the TOC problem under consideration here. A logical extension of this idea is that, as insect metabolomes become available, even HPLC-based study of non-fluorescent primary and secondary metabolite abundance may offer approaches to staging. Second, the underlying biochemistry of insect fluorescence is of interest. Engineered fluorescence for tagging of biological control species has been used by infectious disease entomologists (35) but the use of intrinsic fluorescence characteristics for identification and ecological studies lags behind other fields such as the study of coral larval fluorescence across lifecycle and ecozone (36). In the food storage setting, the intrinsic fluorescence of infesting insect species has been used as a simple and robust indicator of contamination (27). Here, the authors note that the larval spectra of five species (Mediterranean flour moth, Ephestia kuehniella, Zeller (Lepidoptera: Pyralidae); Saw-toothed grain beetle, Oryzaephilus surinamensis, Linnaeus (Coleoptera: Silvanidae); Rice moth, Corcyra cepahlonica, Stainton (Lepidoptera: Pyralidae); Red flour beetle, Tribolium castaneum, Herbst (Coleoptera: Tenebrionidae); and Confused flour beetle, Tribolium confusum, Jacquelin du Val (Coleoptera: Tenebrionidae) all exhibited maximal excitation at ~350nm and emission maxima at ~425 nm. They suggest that larval fluorescence may be attributed to a common fluorophore, perhaps the pterin compounds. These pteridine ring-containing compounds would likely contribute to the fluorescence we observe in the FITC channels, but are by no means the only candidates for the autofluorescence that we observe. Candidate natural auto-fluorophores have been reviewed comprehensively (37). Universal cellular fluorophores in the blue-green region of the spectrum are NADP(H), pterins and flavins (38, 39). Chitin, collagen and elastin are proteins that emit in the blue-green. Orange, red and far-red autofluorescence of the type seen here are less well understood, but may be attributable to uroporphyrinogen and haemoglobin. Both pteridines and lipofuscins have been used to determine age in arthropod taxa (40).

In summary, the present study offers a potential methodology to contribute to TOC determination and aid the death investigator. As a pilot, the study offered new technical and entomological insights but will require significant extension to allow these insights to translate to practice for the forensic scientist.

Acknowledgments

The authors acknowledge discussions with Dr. Madison Lee Goff in the early stages of this project, and his provision of reference samples for C. rufifacies third instar.

Footnotes

*

Supported by the NIH P20MD006084 (HT), NIH 2P20GM103466 (HT) NSF EPSCOR EPS-0903833 (HT), AFRL RCP (FA8650-13-C-5800, HT and DOC).

References

  • 1.Benecke M. A brief history of forensic entomology. Forensic Sci Int. 2001 Aug 15;120(1–2):2–14. doi: 10.1016/s0379-0738(01)00409-1. [DOI] [PubMed] [Google Scholar]
  • 2.Byrd J, Castner J. Forensic entomology: the utility of arthropods in legal investigations. 2. London, U.K: CRC Press; 2009. [Google Scholar]
  • 3.Goff ML, Lord WD. Entomotoxicology. A new area for forensic investigation. Am J Forensic Med Pathol. 1994 Mar;15(1):51–7. [PubMed] [Google Scholar]
  • 4.Joseph I, Mathew DG, Sathyan P, Vargheese G. The use of insects in forensic investigations: An overview on the scope of forensic entomology. J Forensic Dent Sci. 2011 Jul;3(2):89–91. doi: 10.4103/0975-1475.92154. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 5.Oliveira-Costa J, Mello-Patiu C. Application of forensic entomology to estimate of the postmortem interval (PMI) in homocide investigations by the Rio de Janeiro Police Department in Brazil. Internet Journal of Forensic Medicine and Toxicology. 2004;5(1):40–4. [Google Scholar]
  • 6.Tabor K. PhD thesis. Blacksburg, VA: Virginia Polytechnic Institute and State University; 2004. Succession and development of carrion insects of forensic importance. [Google Scholar]
  • 7.Tomberlin JK, Benbow ME. Forensic entomology: international dimensions and frontiers. Boca Raton, FL: CRC Press, Taylor & Francis Group; 2015. [Google Scholar]
  • 8.Catts EP, Goff ML. Forensic entomology in criminal investigations. Annu Rev Entomol. 1992;37(1):253–72. doi: 10.1146/annurev.en.37.010192.001345. [DOI] [PubMed] [Google Scholar]
  • 9.Replogle J, Lord WD, Budowle B, Meinking TL, Taplin D. Identification of host DNA by amplified fragment length polymorphism analysis: preliminary analysis of human crab louse (Anoplura: Pediculidae) excreta. J Med Entomol. 1994 Sep;31(5):686–90. doi: 10.1093/jmedent/31.5.686. [DOI] [PubMed] [Google Scholar]
  • 10.Tomberlin JK, Mohr R, Benbow ME, Tarone AM, VanLaerhoven S. A roadmap for bridging basic and applied research in forensic entomology. Annu Rev Entomol. 2011;56:401–21. doi: 10.1146/annurev-ento-051710-103143. [DOI] [PubMed] [Google Scholar]
  • 11.Boatright SA, Tomberlin JK. Effects of temperature and tissue type on the development of Cochliomyia macellaria (Diptera: Calliphoridae) J Med Entomol. 2010 Sep;47(5):917–23. doi: 10.1603/me09206. [DOI] [PubMed] [Google Scholar]
  • 12.Mohr RM, Tomberlin JK. Environmental factors affecting early carcass attendance by four species of blow flies (Diptera: Calliphoridae) in Texas. J Med Entomol. 2014 May;51(3):702–8. doi: 10.1603/me13149. [DOI] [PubMed] [Google Scholar]
  • 13.Nunez-Vazquez C, Tomberlin JK, Cantu-Sifuentes M, Garcia-Martinez O. Laboratory development and field validation of Phormia regina (Diptera: Calliphoridae) J Med Entomol. 2013 Mar;50(2):252–60. doi: 10.1603/me12114. [DOI] [PubMed] [Google Scholar]
  • 14.Flores M, Longnecker M, Tomberlin JK. Effects of temperature and tissue type on Chrysomya rufifacies (Diptera: Calliphoridae) (Macquart) development. Forensic Sci Int. 2014 Oct 13;245C:24–9. doi: 10.1016/j.forsciint.2014.09.023. [DOI] [PubMed] [Google Scholar]
  • 15.Sharma R, Garg RK, Gaur JR. Various methods for the estimation of the post mortem interval from Calliphoridae: a review. Egyptian J of Forensic Sci. 2013;5(1):1–12. [Google Scholar]
  • 16.Greenberg B, Szyska ML. Immature stages and biology of 15 species of Peruvian calliphoridae (Diptera) Annals of the Entomological Society of America. 1984;77(1):488–517. [Google Scholar]
  • 17.Greenberg B, Tantawi TI. Different developmental strategies in two boreal blow flies (Diptera: Calliphoridae) J Med Entomol. 1993 Mar;30(2):481–4. doi: 10.1093/jmedent/30.2.481. [DOI] [PubMed] [Google Scholar]
  • 18.Liu D, Greenberg B. Immature stages of some flies of forensic importance. Annals of the Entomological Society of America. 1989;82(1):80–93. [Google Scholar]
  • 19.Mendonca PM, Dos Santos-Mallet JR, De Carvalho Queiroz MM. Ultrastructural of larvae and puparia of the blowfly Chrysomya megacephala (Diptera: Calliphoridae) Microsc Res Tech. 2012;75(1):935–9. doi: 10.1002/jemt.22017. [DOI] [PubMed] [Google Scholar]
  • 20.Nwaneshiudu A, Kuschal C, Sakamoto FH, Anderson RR, Schwarzenberger K, Young RC. Introduction to confocal microscopy. J Invest Dermatol. 2012 Dec;132(12):e3. doi: 10.1038/jid.2012.429. [DOI] [PubMed] [Google Scholar]
  • 21.Wilson T. Optical sectioning in fluorescence microscopy. J Microsc. 2011 May;242(2):111–6. doi: 10.1111/j.1365-2818.2010.03457.x. [DOI] [PubMed] [Google Scholar]
  • 22.Niederegger S, Wartenberg N, Spiess R, Mall G. Simple clearing technique as species determination tool in blowfly larvae. Forensic Sci Int. 2011 Mar 20;206(1–3):e96–8. doi: 10.1016/j.forsciint.2011.01.012. [DOI] [PubMed] [Google Scholar]
  • 23.Sukontason K, Methanitikorn R, Sukontason KL, Piangjai S, Olson JK. Clearing technique to examine the cephalopharyngeal skeletons of blow fly larvae. J Vector Ecol. 2004 Jun;29(1):192–5. [PubMed] [Google Scholar]
  • 24.Craemer C, Dippenaar-Schoeman AS, Eardley CD, Grobbelaar E, Mansell MW, Millar IM, et al. Collecting and preserving insects and arachnids. Pretoria, South Africa: National Collections of Insects and Arachnids Biosystematics Division, Australian Research Council Plant Protection Research Institute; 2000. [Google Scholar]
  • 25.Nation JL. Insect physiology and biochemistry. 2. Boca Raton, FL: CRC Press/Taylor & Francis; 2008. [Google Scholar]
  • 26.Sukontason K, Piangjai S, Siriwattanarungsee S, Sukontason KL. Morphology and developmental rate of blowflies Chrysomya megacephala and Chrysomya rufifacies in Thailand: application in forensic entomology. Parasitol Res. 2008 May;102(6):1207–16. doi: 10.1007/s00436-008-0895-6. [DOI] [PubMed] [Google Scholar]
  • 27.Abels JP, Ludescher RD. Native fluorescence from juvenile stages of common food storage insects. J Agric Food Chem. 2003 Jan 29;51(3):544–9. doi: 10.1021/jf020775m. [DOI] [PubMed] [Google Scholar]
  • 28.Samerjai C, Sanit S, Sukontason K, Klong-Klaew T, Kurahashi H, Tomberlin JK, et al. Morphology of puparia of flesh flies in Thailand. Trop Biomed. 2014 Jun;31(2):351–61. [PubMed] [Google Scholar]
  • 29.Buszczak M, Segraves WA. Drosophila metamorphosis: the only way is USP? Current Biology: CB. 1998 Dec 3;8(24):R879–82. doi: 10.1016/s0960-9822(07)00550-7. [DOI] [PubMed] [Google Scholar]
  • 30.Riddiford LM, Cherbas P, Truman JW. Ecdysone receptors and their biological actions. Vitam Horm. 2000;60:1–73. doi: 10.1016/s0083-6729(00)60016-x. [DOI] [PubMed] [Google Scholar]
  • 31.Segraves WA. Steroid receptors and orphan receptors in Drosophila development. Sem Cell Biol. 1994 Apr;5(2):105–13. doi: 10.1006/scel.1994.1014. [DOI] [PubMed] [Google Scholar]
  • 32.Thummel CS. From embryogenesis to metamorphosis: the regulation and function of Drosophila nuclear receptor superfamily members. Cell. 1995 Dec 15;83(6):871–7. doi: 10.1016/0092-8674(95)90203-1. [DOI] [PubMed] [Google Scholar]
  • 33.Sanit S, Sukontason KL, Sribanditmongkol P, Klong-Klaew T, Samerjai C, Sontigun N, et al. Surface ultrastrucure of larva and puparia of blow fly Hypopygiopsis tumrasvini Kurahashi (Diptera: Calliphoridae) Parasitol Res. 2012 Dec;111(6):2235–40. doi: 10.1007/s00436-012-3072-x. [DOI] [PubMed] [Google Scholar]
  • 34.Gabre RM, Adham FK, Chi H. Life table of Chrysomya megacephala (Fabricius) (Diptera: Calliphoridae) Acta Oecologica. 2005;27(3):179–83. [Google Scholar]
  • 35.Harris AF, Nimmo D, McKemey AR, Kelly N, Scaife S, Donnelly CA, et al. Field performance of engineered male mosquitoes. Nat Biotechnol. 2011 Nov;29(11):1034–7. doi: 10.1038/nbt.2019. [DOI] [PubMed] [Google Scholar]
  • 36.Roth MS, Fan TY, Deheyn DD. Life history changes in coral fluorescence and the effects of light intensity on larval physiology and settlement in Seriatopora hystrix. PLoS One. 2013;8(3):e59476. doi: 10.1371/journal.pone.0059476. [DOI] [PMC free article] [PubMed] [Google Scholar]
  • 37.Roshchina V. Vital autofluorescence: application to the study of plant living cells. International Journal of Spectroscopy. 2012;2012 doi: 10.1155/2012/124672. Artcle ID 124672. [DOI] [Google Scholar]
  • 38.Pöhlker C, Huffman JA, Pöschl U. Autofluorescence of atmospheric biosaerosols-fluorescent biomolecules and potential interferences. Atmos Meas Tech. 2011;4(5):5857–933. [Google Scholar]
  • 39.Wolfbeis OS. The fluorescence of organic natural products. In: Schulman SG, editor. Molecular luminescence spectroscopy: methods and applications. New York, NY: John Wiley & Sons; 1985. pp. 167–370. [Google Scholar]
  • 40.Robson SKA, Crozier RH. An evaluation of two biochemical methods of age determination in insects (pteridines and lipofuscins) using the ant Polyrhachis sexpinosa Latrielle (Hymenoptera: Formicidae) Australian Journal of Entomology. 2009;48(2):102–6. [Google Scholar]

RESOURCES