Pectoralis Muscles

Participants were asked to abstain from caffeine-containing food and drink prior to the test and to only consume a light meal 2 h prior to testing. Participant’s skin was cleaned (shaved if necessary) and prepared for the attachment of the ECG electrodes. The electrodes were placed in a CM5 configuration [right fifth interspace, manubrium and left fifth interspace (Dash 2002 )], ensuring that they did not interfere with the fit of the HRM strap (Polar H7). The electrode belt was dampened and placed following Polar’s guidelines, tightly but comfortably just below the chest muscles. Resting measurements were conducted in two positions, supine and, following an active orthostatic challenge, standing in a quiet laboratory, with a temperature of 20.6 ± 1.0 °C. Recordings lasted for 10 min in the supine position and 7 min in standing position. In order to control for the influences of respiration on HRV (Song and Lehrer 2003 (link)) participants matched their breathing frequency to an auditory metronome set at 0.20 Hz (12 breaths min⁻¹). No attempt was made to control the participant’s tidal volume (Pöyhönen et al. 2004 (link)).

Breast density was measured by using fully automated software. Absolute dense area and area percent density (PD %) were estimated by using a publically available software tool [31 ], the Laboratory for Individualized Breast Radiodensity Assessment (LIBRA), based on our previously proposed adaptive multi-cluster fuzzy c-means segmentation algorithm [32 (link)]. The LIBRA algorithm has been previously validated against the current standard semi-automated Cumulus method [33 (link)], showing similar agreement for both raw (i.e., “For Processing”) and vendor post-processed (i.e., “For Presentation”) digital mammograms (Fig. 1) [32 (link)], for the same vendor used in this study. Briefly, the algorithm first applies an edge-detection algorithm to delineate the boundary of the breast and the pectoral muscle. An adaptive multi-class fuzzy c-means algorithm is applied to identify and partition the image gray levels (Fig. 1b) within the mammographic breast tissue area, B_A, into regions (i.e., clusters) of similar x-ray attenuation (Fig. 1c). These clusters are then aggregated by a support-vector machine classifier to a final absolute dense area, D_A, segmentation (Fig. 1d). The ratio of the absolute dense area to the total breast area is used to obtain a measure of breast percent density (PD %): $\mathrm{P}\mathrm{D}\mathrm{\%}=\frac{D_A}{B_A}$
Fig. 1

Example of density segmentation using the LIBRA software tool. a Left mediolateral oblique “For Processing” raw mammogram of a 57-year-old woman with a negative screening exam. b Breast image intensity histogram with fuzzy c-means clustering centroids (vertical lines). c Intensity-clustered breast image. d The final breast and dense tissue segmentation. LIBRA Laboratory for Individualized Breast Radiodensity Assessment

Absolute dense tissue volume and volume percent density (VD %) were automatically assessed by using FDA-cleared software (Quantra™ version 2.0; Hologic Inc., Bedford, MA, USA). The algorithm is based on the widely validated method of Highnam et al. [34 (link)] adapted for digital mammography [35 ]. Briefly, this method quantifies the thickness of dense (i.e., fibroglandular) tissue within each image pixel based on physical parameters of the breast and the imaging system as well as on imaging physics of individual exposures, such as attenuation coefficients for breast tissue, x-ray spectra for the target material, x-ray energy (i.e., peak kilovoltage), exposure, and organ dose (i.e., decigray). Aggregation of the per-pixel volumes for the entire breast allows estimation of the total breast volume, B_V, and dense tissue volume, D_V. The ratio of absolute dense tissue volume to absolute breast volume provides a measure of VD % as: $\mathrm{V}\mathrm{D}\mathrm{\%}=\frac{D_V}{B_V}$
Lastly, for comparison with the automated density measures, we also obtained standard four-category BI-RADS density estimates via retrospective review of archived clinical reports, in which the density assessment was made at the time of routine clinical evaluation by the interpreting breast radiologist for that individual mammography study.

Pathogenesis studies were conducted with specific pathogen free (SPF) white leghorn chickens (Gallus gallus domesticus), broad breasted white turkeys (Meleagris galopova) and Pekin ducks (Anas platyrhynchos domesticus). Chickens were obtained at 2 weeks of age from a commercial supplier of SPF animals (Charles-River SPAFAS, Franklin, CT) and were housed in isolators until they were exposed to the virus at 4 weeks of age. There are no sources of turkeys and ducks which are maintained as SPF or free from viral or bacterial respiratory diseases therefore turkeys and ducks were obtained from commercial hatcheries at hatch and were housed in isolators until they were exposed to virus at 2 weeks of age. All birds were obtained from flocks with no antibody or prior exposure to AI virus. The birds were housed in glove-port isolators (Allentown Caging, Allentown, NJ) with ad libitum access to feed and water before and after exposure to the viruses. Ducks and turkeys were exposed to the virus at 2 weeks instead of 4 weeks of age to accommodate the larger size and faster growth rates of these species in the animal facilities. Since the immune systems of all three species at these ages are considered to be relatively immature, this difference is not expected to impact species associated differences in susceptibility to LPAIV infection and disease. Animals were cared for in accordance with established humane procedures and biosecurity guidelines.
Thirteen to 15 of each species were inoculated with 10⁶ EID₅₀ per bird in 0.1 ml by the intrachoanal route. The birds were monitored daily for clinical disease signs which were scored as follows: 0 = no signs, 1 = mild to moderate respiratory signs (mild depression in ducks), 2 = moderate to severe (i.e. depressed, not eating, neurological signs), 3 = Dead. Oro-pharyngeal and CL swabs were each collected at days 2, 4, 7, 10 and 14 post inoculation (PI) to evaluate virus shed by quantitative real-time RT-PCR (qr-RT-PCR). Three days PI, 3-5 birds from each group were euthanized and necropsied to evaluate gross lesions. Tissues (heart, lung, pancreas/duodemun, kidney, liver, ileum, jejunum, ceca, bursa, thymus, spleen, breast muscle, thigh muscle, brain, nasal cavity, adrenal glands, cecal tonsils, trachea, and reproductive organs) were collected for microscopic evaluation. Serum was collected from ducks at 18 days PI and from chickens and turkeys at 21 days PI to confirm infection status.

All fulmar dissections were performed in
the laboratory following a standard protocol.^{37 ,38} During the dissections, the depth of the subcutaneous fat layer
between the pectoral muscle and the skin was measured at its deepest
with the depth rod of a vernier caliper. For this, the fat tissue
was separated from the muscle tissue and kept attached to the skin
on the side where it was measured. The gastrointestinal tracts (GITs)
were dissected from the esophagus to the anus, along with several
tissue samples for ecotoxicological research (not presented in this
paper). New scalpel blades and gloves were used for each bird, and
the tools were rinsed using soap, Milli-Q water, and ethanol.

This study was conducted with free-living Adélie penguins during the guard stage of chick rearing when adults alternate between taking foraging trips or guarding the young (21st December 2018-14th January 2019) at Dumont d'Urville station, Terre Adélie, East Antarctica (66°40′S; 140°01′E). We captured 58 adults (24 females and 34 males) on the nest, when both adults were attending the nest prior to a changeover. Only one member of a pair was ever sampled, to reduce disturbance time to the nest.
This study took advantage of a blood sample protocol required for a doubly labelled water (DLW) experiment that necessitated two initial blood samples, one taken after capture and a second one taken after a calibration period in which the animals must be held in captivity (Hicks et al., 2020 (link)). Specifically, upon capture, individuals were blood sampled within 3 min of first handling from the tarsus vein. This sample served as a background sample providing the unstressed situation. After blood sampling, birds were weighed to the nearest gram, flipper measured to the nearest millimetre, and injected with 0.3 ml of DLW per kg of body weight into the pectoral muscle [see Hicks et al. (2020) (link) for details]. For future identification, birds were marked with a unique identifying code printed on a piece of marine tape rolled around their back feathers. This entire handling period took about 15 min (study mean) and several previous studies have shown that 15 min of handling evokes a stress response as measured by increases in corticosterone in this species (Cockrem, 2013 (link); Cockrem et al., 2006 (link), 2008 (link)) and also in this population (Marciau et al. under review ). After this handling period, birds were placed in a contained area outside the lab for the DLW to equilibrate (for between 1.6 and 2.7 h). The contained area measured 2×2.80×5.20 m, birds were always placed in the contained area with another penguin to calm them and the space was filled with a layer of snow for comfort and cooling. A second blood sample was taken after the equilibrium period. This sample served as our handling time sample. Hold duration was calculated as the time between the first capture and the second blood sample. Whole blood was kept on ice in Eppendorf tubes for up to 10 min before being centrifuged (10,000 rpm, 10 min) and plasma stored at −80°C until analysed. The molecular sexing of all individuals was carried out at the service Analyses Biologiques of the Centre d'Etudes Biologiques de Chizé (CEBC). DNA extraction was conducted with 2 µl of pellet (red blood cells) and using a chelex resin (Chelex 100 Molecular Biology Resin, Bio-Rad; 10%) associated with Proteinase K (PK) as written in the manufacturer's instructions. We then performed a polymerase chain reaction (PCR) with amplification of the CHD gene following a standard procedure validated on penguins (Lee et al., 2010 (link)).