Oviposition

A natural analytical framework for considering the effects of temperature on malaria transmission intensity is provided by deterministic models for the disease's basic reproductive number, R ₀ , defined formally as the expected number of new cases arising in a naive population after one generation of the parasite from the introduction of a single infectious person [39 -41 ]. These models parameterise malaria transmission in terms of characteristics of, and interactions between, human, vector, and parasite populations [42 -46 (link)]. Those aspects of the transmission cycle affected by temperature are encapsulated in a component of R ₀ known as vectorial capacity [47 (link)], V , which defines the total number of subsequent infectious bites arising from a single person-day of exposure and is classically expressed as:
where m is the number of mosquitoes per human, a is the human feeding rate, p is the daily vector survival rate, and n is the time required for sporogony, the maturation of parasites ingested by mosquitoes during human blood meals into the sporozoite life cycle stage infectious to humans. Expressing vector survival in terms of daily death rate, g where, g = -ln p, and holding constant the rate of adult mosquito recruitment, λ, relative to the human population so that, m = λ/g, vectorial capacity can be rewritten [48 (link)] as:
Temperature can influence all of the terms in this equation. Temperature affects feeding rates, a for example, via effects on vector activity and blood meal digestion [49 -51 ]. Larval ecology and, thus, adult recruitment, λ, are affected by temperatures found in aquatic habitats which play a role in modulating larval development rates and survival [33 (link),34 (link),52 (link),53 ]. Other work has demonstrated how these factors alone can impose limits on habitat suitability for particular anopheline species [53 ]. Adult mosquito recruitment is, however, also driven by a myriad of other climatic and local environmental factors, in particular those associated with the often transitory availability of aquatic oviposition sites. Here we focus on the more pronounced and directly measurable effects of temperature on vectorial capacity: the interaction between vector lifespan, determined by, g and the duration of sporogony, a. Holding a and λ constant, then, we can modify equation (2) to obtain an expression as a function of temperature, T:
Since a and λ are unknown, vectorial capacity cannot be evaluated directly, so we define instead an index of temperature suitability Z(T) that is linearly proportional to V(T) and therefore sufficient for exploring the relative, rather than absolute, effect of temperature on vectorial capacity and, thus, on R ₀ . The index Z(T) can be interpreted as a relative measure of the number of infectious mosquitoes supported in an environment with temperature T, given a constant emergence rate λ. All other things being equal, an environment with, say Z(T), a value of 100 would support twice the vectorial capacity or, equivalently, require half as many vectors to support the same vectorial capacity as one with a Z(T) value of 50. Locations in which Z(T) is zero indicate that no vectors survive long enough to accumulate sufficient degree days for sporogony.

All infections occurred at 22 °C. Either six female Lb17 or five female Lh14 wasps were placed in 35-mm petri dishes filled with 2 ml of fly food and 50 fly larvae (in pilot experiments using D. melanogaster, we found that these wasp/host ratios yielded an approximately equal number of wasp eggs per fly larva). Wasps were removed 2 d later. The proportions of wasps that emerged from each plate were scored and averaged across multiple plates. The average number of flies infected per treatment was approximately 250. While the length of contact between wasps and fly larvae ensures that wasps will have access to host larval stages ideal for successful infection, repeated ovipositions in single fly larvae could result in inflated parasitization success [2 ]. For wasp/fly species pairs that yielded very few wasp offspring, we did not attempt to distinguish refusal to lay eggs, wasp egg developmental arrest, or host immune response against the eggs as potential reasons for lack of wasp success.

Insects used for transcriptome were R. prolixus from a colony kept at UFRJ (Rio de Janeiro), fed with rabbit blood, and raised at 28°C and 70% relative humidity. Adult females (five from each condition) receiving their second blood meal after the imaginal molt were dissected before feeding, twelve hours, twenty-four hours, two days, and five days after blood meal. A group of males (blood fed, five days after blood meal) was dissected to obtain testes. Organs (AM, PM, RE, FB, OV, MT, and TE) were dissected, homogenized in TriZol reagent (Invitrogen, San Diego, CA, USA), and processed as described below. To obtain a whole body (WB) library, nymphs and adults in several stages of feeding plus eggs were collected and extracted with TriZol, as follows: Eggs were collected at the day of oviposition and at days 2, 5 and 7 of development. First instars were collected at fasting (2 weeks after emergence) and at 2, 5 and 7 days after feeding (DAF); second and third instars were collected at fasting and at 2, 5, 7 and 9 DAF. Fourth instars were collected at fasting and at 2, 5, 7, 9 and 12 DAF. Fifth instars were collected at fasting and at 2, 5, 7, 9, 12, 14, 17 and 19 DAF. Adult males and females were collected at fasting and at 2, 5, 7, 9 and 12 DAF. All these 45 RNA preparations were pooled and used to obtain WB cDNA as described below.

The post-attachment period spanned the initial 8 days following tick attachment (day 0 to day 8). During this time, deer were housed individually in pens (Additional file 5: Figure S3) and checked daily to ensure adequate health and wellbeing and to ensure that the capsules remained firmly attached. At day 6 and day 8 post-attachment, the deer were anesthetized in the previously described manner, and the capsules were opened to monitor the condition of the ticks. These time points were selected because I. scapularis was the primary species of concern and reportedly takes approximately 6–11 days to reach engorgement and detach [45 ]. A minimum of 48 h was required between sedations for each deer because of PSU IACUC policy forbidding sedation of animals on consecutive days.
Ticks were easily visible by the naked eye. The inside of each capsule was carefully scanned for attached and detached ticks. The numbers of total ticks recovered and their attachment status (attached, detached), feeding status (flat, partially engorged, fully engorged) and condition (alive, dead) were recorded. Any dead ticks (attached, detached) were removed from deer. At the conclusion of tick observations on day 8 post-attachment, capsules were completely removed, and any live or dead ticks were manually removed from the deer.
Fully engorged, live detached female ticks were collected, weighed to the nearest 0.0001 g using an analytical balance (Mettler-Toledo, LLC, Columbus, OH, USA), and maintained individually in vials. Engorged females were maintained in a desiccator (> 90% relative humidity) and were allowed approximately 14–28 days to complete oviposition [45 ]. After oviposition was completed, females were removed, and egg masses weighed to the nearest 0.0001 g. Egg masses were monitored for the emergence of larvae, with eggs embryonating within approximately 35–50 days [45 ]. Egg masses were monitored for approximately 2–3 weeks to estimate the proportion of hatched eggs.

At the age of 70 weeks, a total of 120 hens were randomly selected to evaluate dietary interventions of body phosphorus rhythms. The hens were fed with 4 phosphorus regimens: (1) RR, provided with regular phosphorus diet at both 09:00 and 17:00 (conventional feeding without considering daily rhythms of body phosphorus metabolism); (2) RL, provided with regular phosphorus diet at 09:00 and low phosphorus diet at 17:00 (dynamic feeding converse to the body phosphorus rhythms found in Exp. 1); (3) LR, provided with low phosphorus diet at 09:00 and regular phosphorus diet at 17:00 (dynamic feeding consistent with the body phosphorus rhythms found in Exp. 1); (4) LL, provided with low phosphorus diet at both 09:00 and 17:00 (direct restriction without considering daily rhythms of body phosphorus metabolism). Each feeding regimens included 6 replicates, and each replicate contained 5 hens. The regular and the low phosphorus diet contained 0.32% and 0.14% NPP, respectively. The feeding trial lasted for 12 weeks (according to the literature, changes in eggshell and bone mineralization status could be observed in 8 to 12 weeks after dietary phosphorus interventions in laying hens) [26 (link), 27 (link)]. On the last 3 d of the feeding trial, all the eggs were collected for egg quality analysis. On the last day of the feeding trial, two egg-laying hens were randomly selected from each replicate (sampled at 6 and 18 h post-oviposition, respectively). The following samples were collected: blood (for serum), uterine (stored at −80 ℃, for Western-blotting analysis ), femur (left side, stored in 4% paraformaldehyde for histological analysis; right side, stored at −80 ℃ for the determination of mineralization status and gene expressions).

Hy-Line Brown laying hens were fed with a regular diet (corn-soybean meal-based; containing 0.32% non-phytate phosphorus (NPP); Table 1) start from 35 weeks of age. On the last day of age 40 weeks, a total of 60 hens that laid eggs between 07:30−08:30 were randomly selected to evaluate the daily phosphorus rhythms. Of them, 45 hens were euthanized for sample collection, and the other 15 hens were used to study the feed intake and calcium/phosphorus excretion rhythms. For sample collection, the 45 hens were sampled according the oviposition cycle: at oviposition, at 6, 12, 18 h post-oviposition, and at the next oviposition, respectively, with 9 hens sampled at each of the time point. The following samples were collected: blood (for serum), uterine (stored at −80 ℃, for Western-blotting analysis), femur (in 4% paraformaldehyde, for histological analysis) and kidney (stored at −80 ℃, for Western-blotting analysis). For the other 15 hens, the feed intake was recoded and the excreta was collected at the following intervals: from oviposition to 6 h post-oviposition, from 7 to 12 h post-oviposition, from 13 to 18 h post-oviposition, from 19 h post-oviposition to the next oviposition.Table 1

Composition and nutrient concentrations of basal diet (%, unless noted, as-is basis)

Item	Low phosphorus	Regular phosphorus
Ingredients
Corn	56.69	56.69
Soybean meal	25.77	25.77
Distillers dried grains with solubles	4.00	4.00
Calcium carbonate	9.73	9.04
Dicalcium phosphate	-	1.15
Soybean oil	1.51	1.51
Sodium chloride	0.26	0.26
DL-Methionine	0.18	0.18
Choline chloride	0.15	0.15
Montmorillonite	0.71	0.25
Premix1	1	1
In total	100.00	100.00
Nutrient levels
Metabolizable energy, kcal/kg (calculated)	2,600	2,600
Crude protein (calculated)	16.5	16.5
Total phosphorus (calculated/analyzed)	0.34/0.34	0.53/0.49
Non-phytate phosphorus (calculated)	0.14	0.32
Calcium (calculated/analyzed)	3.50/3.47	3.50/3.52

¹Provided per kilogram of diet: manganese 60 mg, copper 8 mg, zinc 80 mg, iodine 0.35 mg, selenium 0.3 mg, vitamin A 8000 IU, vitamin E 30 mg, vitamin K₃ 1.5 mg, thiamine 4 mg, riboflavin 13 mg, pantothenic acid 15 mg, nicotinamide 20 mg, pyridoxine 6 mg, biotin 0.15 mg, folic acid 1.5 mg, and cobalamin 0.02 mg