Interpretation of the morphological adaptations associated with viviparity in the Tsetse fly (Glossina morsitans) by three dimensional analysis

Tsetse flies (genus Glossina), the sole vectors of African trypanosomiasis, are distinct from other disease vectors, and most other insects, due to dramatic morphological and physiological adaptations required to support their unique life histories. These evolutionary adaptations are driven by demands associated with their strict dietary and reproductive requirements. Tsetse reproduce by obligate viviparity which entails obligate intrauterine larval development and provisioning of nutrients for the developing larvae. Viviparous reproduction reduces reproductive capacity/rate which also drives increased inter- and intra-sexual competition. Here, we use phase contrast microcomputed tomography (pcMicroCT) to perform a three-dimensional (3D) analysis of viviparity associated morphological adaptations of tsetse female reproductive tract. These include 1) abdominal modifications facilitating the extreme abdominal distention required during blood feeding and pregnancy; 2) abdominal and uterine musculature required for parturition of developed larvae; 3) reduction of ovarian structure and capacity; 4) structural features of the spermatophore form in the female uterus to enhance semen/sperm delivery and inhibition of insemination by competing males; 5) uterine morphological features facilitating expansion and contraction before, during and after pregnancy; 6) milk gland structural optimizations facilitating nutrient incorporation and transfer into the uterus. The use of pcMicroCT provides unprecedented opportunities for examination and discovery of internal morphological features not possible with traditional microscopy techniques and new opportunities for comparative morphological analyses over time and between species.


Introduction
Tsetse flies (genus Glossina) are the sole vectors of African trypanosomes, the causative agents of Sleeping Sickness in humans and Nagana animals. These are fatal diseases predominantly affecting marginalized populations in sub-Saharan Africa which cause severe health and economic impacts in affected countries 1 . Vector control methods are a primary method of control as tsetse flies have a low reproductive capacity relative to other insects 2 . This is due to specializations in tsetse's reproductive biology that results in the production of a small number of offspring over a long period of time. Tsetse females reproduce by obligate viviparity which is defined as having intrauterine larval development and provision of all larval nourishment for the duration of its development [3][4][5] . Each gonotrophic cycle lasts 10 days, which restricts progeny per female to 8-10 during their lifespan.
Mating and transfer of seminal fluid into the uterus stimulates behavioral and physiological changes in female tsetse flies. Within 48 hours of mating, females become refractory to additional mating attempts by other males, start taking larger blood meals and begin to ovulate. This physiological response is exploited for control purposes via use of sterile insect technique 6,7 . However, in Glossina, little is known regarding the biology underlying the post mating response.
How the post mating response in Glossina is triggered by mating associated stimuli and what happens to internal female morphology in response to these stimuli remains unknown. The reproductive tissues of tsetse flies are highly derived relative to that of other dipterans. The most dramatic of the reproductive modifications include the reduction of ovarian capacity 9 , enhanced uterine musculature 10 and modification of the female accessory gland into a tubular ramified milk-producing organ [11][12][13] . Descriptions of Glossina reproductive tissues have been made via a variety of media including detailed hand drawn illustrations of dissected tissues as well as via various light and electron based microscopy techniques 5,11,[13][14][15][16][17][18][19][20] . Most of these techniques require compromises in one form or another which can limit the scope of the analysis.
Most microscopy techniques are destructive to different extents in that they require the sample be dissected and/or cut into thin sections for staining and imaging. The process of dissection, fixation and sectioning can cause mechanical disruption of tissue shape and positioning within the insect. These processes can also damage delicate features and interactions within and between tissues. Most microscopy-based imaging generates two-dimensional images. The lack of the third dimension confines the context and scope of the information contained within the image and can limit its informative capacity.
While some light microscopy-based techniques such as confocal laser scanning microscopy have the capability to generate stacked three-dimensional (3D) images.
Howevver, this technique is restricted by sample section thickness/opacity which limits the potential sample thickness to ~20-50 µm and is not applicable at the level of whole mounted tissues 20 .
X-ray Micro Computed Tomography (MicroCT) provides an alternative that addresses many of the described issues 21 . This technique uses an X-ray beam to virtually section through a sample across 180° of rotation resulting in an image stack representing the entire volume of the specimen. The high energy of X-rays allows biological samples of any thickness to be visualized and does not result in structural damage to the sample, which allows for repeated scans or further use of the sample in alternative visualization techniques. The primary challenge of using this technique is generation of contrast in soft tissues associated with biological samples so that features of interest can be visualized in detail. With traditional microCT this requires sample fixation followed by staining with a metallic agent such as iodine 22 . The iodine functions to absorb energy from the X-ray beam to boost contrast and define soft tissue features by absorbance.
Alternatively, the use of a system where the energy of the X-ray beam can be adjusted to provide optimum contrast is ideal. These conditions are available at synchrotronbased facilities which allow for tuning of the X-ray beam energy. In addition, synchrotron based microCT allows for imaging via differential phase contrast rather than traditional absorbance. Phase contrast microCT (pcMicroCT) facilitates visualization of phase changes in the X-ray beam resulting from its passage through materials with different refractive indices. This allows for high contrast 3D visualizations of sample composition with or without metallic staining 23 .
Use of microCT in the analyses of female reproductive tissues in Drosophila melanogaster revealed that mating induces morphological changes in female reproductive tissues over the course of the post mating transition 24 . These changes include looping/unlooping of the uterus and oviduct, repositioning of reproductive tract within the abdomen and mating induced tissue damage resulting from the male reproductive organs. Its hypothesized that the tissue damage may allow male seminal secretions access to the hemocoel of the female. Post mating morphological changes are also observed in the oviduct which connects the ovaries to the uterus. Prior to mating, the lumen of the oviduct is closed and appears to be a solid cylindrical tube of muscle and epithelial cells. Following mating the oviduct undergoes a developmental program resulting in the formation and opening of the duct lumen facilitating the passage of oocytes from the ovaries into the uterus 25 .
In this work, the abdominal tissues from whole freshly mated female tsetse flies are imaged via pcMicroCT to evaluate the capacity of this technique to perform 3D analyses of tsetse reproductive morphology and for future comparative analyses of post-mating changes. Analysis of the resulting data has provided detailed 3D visualizations of external and internal morphological features and provides new insights into the functional roles of structural features of Glossina reproductive organs.

Biological materials
Tsetse flies (Glossina morsitans) utilized in this analysis were obtained as pupae from the colony maintained at the Institute of Zoology at the Slovak Academy of Sciences in Bratislava, Slovakia. Flies were reared in the Tupper Hall arthropod containment level 2 insectary in the UC Davis School of Veterinary Medicine. Flies are maintained in an environmental chamber at 25ºC and 75% relative humidity with 12:12 light/dark photoperiod. Flies receive defibrinated bovine blood meals via an artificial feeding system Mondays, Wednesdays and Fridays as described 26 . Sterile defibrinated bovine blood for feeding is obtained from Hemostat Laboratories (Dixon, CA).

Sample collection
Tsetse pupae were placed into an eclosion cage and monitored for teneral females daily. Teneral flies were anesthetized on ice and sorted into cages by sex. At five days post eclosion, individual virgin females were combined with males in mating cages.
Cages were observed for mate pairing. If pairing was not observed within 10 minutes the male was removed from the cage and a new male introduced. Tsetse flies require at least one hour of pairing for completion of the transfer and formation of the spermatophore 27 . Mate pairs lasting for less than 60 minutes were removed from the sample pool. Within an hour of mating completion, females were anesthetized on ice for sample preparation.

Sample preparation, fixation and staining
Chilled flies were prepared for fixation by removal of the legs and wings to allow the fixative to permeate into the haemocoel. The fly was then placed into Bouins fixative solution (acetic acid 5%, formaldehyde 9% and picric acid 0.9%) and incubated overnight at room temperature. Following fixation, flies were dehydrated using a graded series of ethanol washes (10%, 30%, 50%, 70% and 95%). Each wash was performed for 1 hour at room temperature. Flies were then stained in 1% iodine in 100% ethanol for 24 hours. After staining, flies were washed 3 times in 100% EtOH for 30 mins per wash.

Phase contrast micro computed tomography
During imaging, samples must remain in a fixed position with no movement during the scanning process to ensure proper alignment of the image stack. In preparation for imaging, fixed and stained flies were transferred into a 1.5 mL Eppendorf tube containing unscented Purell hand sanitizer (Gojo Industries, Akron OH). The specimen was gently pushed down to the bottom of the sample tube using a pipette tip. To ensure samples remained immobilized during scanning, the bottom of another 1.5 mL Eppendorf tube was cut off and pushed into the sample tube for use as a wedge. The wedge was gently pushed down into the sample tube until the specimen was secured between the wall of the container and the wedge. Once the specimen was secured, the remaining volume of the sample tube was filled with Purell. The sample tube was modified to attach to the sample holder (chuck) in the MicroCT imaging hutch by hot Software available at http://www.theobjects.com/dragonfly. Image volumes were imported into Dragonfly and down sampled to 16-bit depth at half resolution (1280x1280) to improve computer performance during analysis and segmentation. The image volume was cropped to eliminate portions of the volume not occupied by the sample. Upon import, dataset contrast and sharpness were enhanced using the Unsharp image processing function with a kernel size of 7, standard deviation of 3 and unsharp factor of 3. Tissue segmentation and region of interest definitions were performed using a combination of algorithmic and manual methods. Images were captured and exported from within Dragonfly. Tissue volumes, surface areas and thickness mapping were calculated by measurement of segmented voxels by Dragonfly with each voxel representing 1.6 uM 3 . The associated video was generated using the Dragonfly movie maker function. The video is available for download from https://drive.google.com/file/d/1bHU2A6Fsxb_ZuJkg3gnbfAWXuQmr-TLG/view?usp=sharing.

Abdominal structural and cuticular adaptations for blood feeding and pregnancy
The analysis described here encompasses the analysis of the last four abdominal segments of a female tsetse fly (Glossina morsitans) immediately after copulation ( Figure 1). This region contains the entirety of the reproductive tract as well as tissues from other organ systems such as the digestive tract, abdominal musculature, respiratory system and the fat body (nutrient storage/metabolism). The presence of these other systems within the scan provides spatial context of how these tissues interact within other abdominal tissues. It also provides clues to the structural adaptations required to accommodate the massive changes in abdominal volume required during blood feeding and pregnancy. As in most blood feeding insects, tsetse flies ingest large volumes of blood relative to their body size. After mating, females on average take blood meals weighing between 50-60 mg 30 . In addition, pregnant females accommodate fully developed 3 rd instar larvae equivalent in mass to themselves. Newly eclosed female flies weigh ~18 mg and just after a blood meal a pregnant female can weigh over 90 mg prior to water elimination via diuresis 30 . Thus a 5-fold change in mass occurs over the course of a pregnancy cycle. It follows that to accommodate these dynamic changes, abdominal volume can increase by up to 60%.
A detailed view of exterior ( Figure 1A+B) and interior ( Figure 1C+D Ovarian reduction and oviduct folding reduce constraints on blood meal volume and larval size (Figure 4) Another major adaptation to Glossina reproductive physiology, relative to oviparous Diptera, is the reduction in ovarian capacity. The ovaries of most oviparous Diptera contain dozens of ovarioles with some or all containing vitellogenic oocytes at the same time 33 . In Glossina, each ovary contains two ovarioles with only one mature oocyte completing development per gonotrophic cycle 8,34,35 (Figure 4A+B). Cross section through the developing ovarian follicles shows the fully developed primary follicle with the tertiary ovarian follicle in close association inside the right ovary ( Figure 4C). In the primary follicle, the oocyte has filled with yolk protein crystals and lipids and the nurse cells have emptied into the oocyte and collapsed. This oocyte has completed most of its development and is almost ready for ovulation into the uterus.
Tsetse flies require mating associated stimuli to undergo ovulation [36][37][38]  During ovulation, the fully developed oocyte moves out of the ovary, through the oviduct and into the uterus. In Glossina, the oviduct and its associated tissues form a structure called the oviductal shelf which is a sleeve of tissue connecting the ovaries to the uterus ( Figure 4D). The oviductal shelf folds over on itself and occupies most of the dorsal surface of the uterus. The lack of connective musculature on the dorsal anterior portion of the uterus is likely due to the spatial constraints associated with these structures. The folded conformation of this tissue results in the oviduct forming a hairpin turn prior to its opening into the uterus. The compacted structure of the oviductal shelf in addition to reduced ovarian capacity/production optimizes the space occupied by the reproductive tract within the abdominal cavity. These optimizations allow for larger blood meal volumes and increased nutrient storage capacity by the fat body between larvigenic cycles. During ovulation, the oviduct and oviductal shelf likely must expand and straighten to allow the oocyte to pass into the uterus. Whether the oviductal shelf returns to this conformation after the first ovulation and during pregnancy will need to be evaluated in scans taken later in the gonotrophic cycle.
The sagittal section of the uterus reveals that the anterior wall of the uterus also appears folded upon itself and is likely capable of expanding to accommodate the large volume required by the developing intrauterine larvae ( Figure 4D). In addition, the tissue constituting the anterior and posterior uterine walls shows patterning which may represent additional folding that provides elasticity and flexibility to the uterine walls.
The entire reproductive tract is optimized to conserve space while providing the capacity to expand and occupy most of the abdominal volume during intrauterine larval development when the larva is equivalent in mass to the mother.
The spermatophore facilitates sperm storage within the spermatheca and acts as a physical barrier to insemination by competing males (Figure 5).
In Glossina, physical and chemical mating stimuli are required to initiate ovulation and activate other post mating changes including initiation of sexual refractoriness, accelerated oocyte development, increased host seeking/blood feeding and ingestion of increased blood meal volumes 27,34,36,39,40 . Mating involves the transfer of a large volume of male accessory gland derived proteins and biochemicals which form a well-defined structure called a spermatophore in the uterus of the female [41][42][43][44] . The center of this structure is hollow and contains sperm to be stored in the spermathecal organ of the female. The outer wall of this structure is composed of two layers with differing ultrastructural characteristics and has an opening at the dorsal anterior 45 . Visualization of the spermatophore reveals that the outer walls are molded to the interior of the uterus and that it occupies a significant volume of intrauterine space ( Figure 5B). However, the spermatophore resulting from the second mating attempt is usually stuck to the back of the first which results in a physical barrier preventing sperm from exiting the lacuna of the spermatophore. Even if the sperm were able to exit the spermatophore it is unlikely they would be able to navigate around the first spermatophore to access the spermathecal ducts.
Among the Diptera, the use of spermatophores by males is infrequent 46 . However, in tsetse flies inter-and intrasexual competition is intense due to the low reproductive rate of females. There is evidence that there is cryptic selection by females as mating with substandard males can result in poor sperm uptake and continued receptivity to other males 47 . Comparative analysis of male seminal protein genes between six Glossina species revealed them to be the most rapidly evolving genes in the genome with observed differences in gene number and sequence variability 48 . The use of the spermatophore by males may increase the probability that a mating attempt is successful in the face of female selective pressures. The structural advantages of the spermatophore for males are that it is difficult for the female to expel, it guides the sperm to their destination and inhibits insemination by sperm from competing males.
Females normally dissolve the spermatophore and expel its remnants ~24 hours post mating and become refractory to further matings ~48 hours post mating 41 .

The milk gland organ maximizes nutrient incorporation via extensive ramification for increased surface area and intimate contact with fat storage tissues (Figure 6)
The final component of the reproductive tract is the milk gland. The milk gland is a heavily modified female accessory gland which is responsible for the synthesis and secretion of a protein-and lipid-rich milk like secretion. The milk gland is a dynamic organ which undergoes cyclical changes in volume throughout the reproductive cycle in correlation with milk production activity 45,46 . In Glossina austeni, it is estimated that the milk gland produces 25 mg of milk secretion per gonotrophic cycle which is equivalent to the weight of an unfed adult female 46,47 . To accomplish this feat the gland must incorporate large amounts of stored lipids and free amino acids from dietary sources 49,50 . The milk gland does not seem capable of protein uptake from the hemolymph and instead imports amino acids which are utilized by the massive arrays of rough endoplasmic reticulum for milk protein synthesis [43][44][45] . The milk gland also harbors the extracellular form of the Glossina obligate symbiont Wigglesworthia glossinidius. The bacteria live in the lumen of the gland and are transferred to the intrauterine larva during lactation [44][45][46] .
Prior analysis of the milk gland has been primarily by 2D microscopic analysis with hand drawn interpretations of the 3D structure of this complex organ. The analysis of the abdominal volume from the pcMicroCT scan has provided a detailed virtual representation of the milk gland which clearly demonstrates the intricate nature of its branching structure ( Figure 6A). The gland consists of three regions, the distal milk gland, the proximal milk gland and the common collection duct. The milk gland is connected to the dorsal surface of the uterus via the common collection duct. This duct connects to the uterus postero-ventrally to the oviductal shelf and just posterior to the opening of the spermathecal duct ( Figure 6B). The duct extends through the musculature of the uterine wall and travels along the right side of the oviductal shelf where it crosses dorsally over the right spermathecal duct. The common collection duct is formed of two cuticle lined tubes lacking in secretory cells that are bundled together by a spiraling musculature, which likely regulates the flow of the milk secretions [47][48][49] . At the point where the ovaries begin, the common collection duct branches into four thick tubules. Two of these cross dorsally over of the spermathecal ducts and spermathecae to expand into the left side of the abdomen while the other two proximal tubules extend to the right. The proximal milk gland lacks the musculature that wraps around the common collecting duct and contains secretory cells that contribute to milk production. An inherent aspect of the tubular structure of the milk gland is that it has a very high surface area to volume ratio. Analysis of the milk gland in the context of the surrounding abdominal tissues demonstrates that it is intimately associated with fat body cells which store large volumes of lipids for milk production ( Figure 6D to this technique is that scanned volumes can be made publicly available similarly to datasets from other high throughput technologies. Users can download these volumes for independent analysis of features that were not a focus of the original analysis.
Finally, the 3D nature of this data allows export of it to new 3D visualization technologies such as virtual reality applications. With the aid of a virtual reality headset, the user can explore the physiology interactively in 3D at an otherwise impossible scale.
The addition of spatial cues with rich visuals leverages the natural capabilities of the human brain to rapidly interpret and remember information provided in this format [52][53][54] . A powerful feature of traditional microscopy is the ability to visualize gene expression patterns or protein localization via techniques such as in situ and immunohistochemical staining. Protocols for performing these types of analyses via microCT are in their infancy and are complicated by accessibility and even diffusion of staining and washing reagents in a whole mounted specimen. In addition, traditional staining reagents do not absorb high energy X-rays making them invisible to this imaging technique. However, metallic staining reagents coupled to horse radish peroxidase reactive substrates show promise as a way to address these issues 55 .
Finally, efficient analysis of the large volumes of data generated by these scans is hindered by the time required for accurate annotation. Segmentation of soft tissues with similar x-ray absorbances is difficult if not impossible for automated algorithms. Phase contrast helps with this but may still be inadequate for basic contrast-based segmentation programs. The tissue segmentations performed for this volume were done mostly by hand with aid from contrast-based analysis tools and predictive algorithms. The results often require a human eye to correctly determine boundaries between adjoining tissues. New methods based on cutting edge artificial intelligence techniques such as machine learning and deep learning are in development. The software used for this analysis Dragonfly version 4.1 for Windows implements deep learning algorithms that utilize manually segmented datasets to train the algorithm which can then be used to accurately segment raw datasets with minimal human intervention. These methods are still in early stages, but will develop rapidly as more data becomes available, computing power increases and more people develop training sets 56

. This technique was recently used to visualize the interactions between
Cordyceps fungus interactions with the brain and musculature of carpenter ants 57