**4.** *Philornis downsi* **larval development in the wild and in the laboratory**

#### **4.1** *Philornis downsi* **larval instars**

*Philornis downsi* larval development is split into three instar development stages. 1st instar larvae generally reside in the naris and ear canals of developing nestlings, but some have also been found moving freely within the nesting material [21, 52, 53]. First instars are commonly collected from 2 to 3 day old nestlings [43]. Late 2nd and 3rd instar larvae are generally free-living, residing within the base of the nest and feeding externally on nestlings at night [14, 45, 46]. These later instar larvae feed

#### **Figure 1.**

*Three larval stages of Philornis downsi. (1) First instar: (A) posterior spiracles, (B) lateral view, (C) ventral view. (2) second instar: (D) posterior spiracles, (E) lateral view, (F) ventral view. (3) third instar: (G) posterior spiracles, (H) lateral view, (I) ventral view. Obtained by the authors from larvae collected on Floreana Island, Galápagos, Ecuador between 2010 and 2014. The photographs were taken using a visionary digital LK imaging system (dun, Inc) with a canon EOS 5DsR camera and capture one pro 11.3.1, phase one (Flinders University). Images were produced using Zerene stacker 1.04, Zerene systems LLC, software, and cropped and resized in Photoshop CS5.*

on the blood and fluids of their host by penetrating the skin of the nestlings [2, 30]. Larval instar morphological descriptions are given by Fessl et al. [44]. The most distinct character between the instars is the posterior spiracles, which change in colour, shape and number of spiracular slits present throughout larval development [44].

**Figure 1(1A)** shows the posterior spiracles of a 1st instar *P. downsi* larva, characterised by their light pigmentation and two oval slits present [44]. The spiracles of a

**57**

*Taxonomic Shifts in* Philornis *Larval Behaviour and Rapid Changes in* Philornis downsi*…*

1st instar larva are separated by slightly more than their diameter. First instar larvae lack anterior spiracles (**Figure 1(1B)**). The posterior spiracles of a 2nd instar larva are similarly round with two oval slits; however, the distance between them is two to three times of their diameter (**Figure 1(2D)**; [44]). Anterior spiracles are present during the 2nd instar, and semicircular in shape, lightly pigmented and visible in (**Figure 1(2E)**). 3rd instar posterior spiracular plates are darkly pigmented and round in shape, distinct C-shaped spiracular slits radiate from median ecdysial scar (**Figure 1(3G)**). Pigmentation of the median ecdysial scar is light in early 3rd instar larvae and becoming darkly pigmented later in the stage. Semi-circular anterior spiracles are retained in 3rd instar larvae (**Figure 1(3H)**). Cephaloskeleton morphology differs between instars as outlined in Fessl et al. [44]. Recent studies report a decrease in *P. downsi* puparia size across 2004–2014 [54]. Common et al. (unpublished data), and hence body size is certainly not a useful method to classify instars. In general, it is recommended to use a suite of morphological characters, including anterior and posterior spiracular morphology, to determine the larval instar.

The developmental period of *Philornis* larvae is associated with the species' larval feeding habit. For example, time to pupation in coprophagous species takes up to 29 days, but only 4–8 days in subcutaneous species [2, 55]. Larval development periods in free-living species such as *P. downsi* are difficult to determine in the wild as the host nest needs to be dismantled to observe the larvae. Early studies of abandoned or failed nests found 1st instar larvae in nests with 1–3 day old nestlings, 2nd instars in nests with 3–6 day old nestlings and 3rd instars in nests with 3–10 day old nestlings [44]. Larval collections following the cessation of activity at host nests

Compared with larval development times in the wild, larval development times under laboratory conditions are longer. First attempts to rear *P. downsi* larvae in the absence of a living host had a low success rate, with only three larvae out of 477 reaching the adult stage after a 36 day development time (mean 18 day larval development, 12 day pupation) [56]. As the diet for rearing larvae in captivity was refined, the success rate increased to 10% and larval development time decreased [57]. Development time in the laboratory ranged from 9 to 10 days from larva to pupa [57] with even faster development times occurring as the rearing conditions have improved [pers. comm. P. Lahuatte]. Egg hatch rates in captivity have been high (96%), with most mortality in 1st instar larvae (77%) [57]. Laboratory-based diets that have been developed in the absence of a bird host are primarily based on chicken blood, with more successful diets including hydrolysed protein and vitamin fortification [57]. The lack of keratin in the diet may be causing elevated 1st instar mortality, as 1st instars consume the keratin of the beak in which they reside [44],

The behaviour of adult *P. downsi* is much less understood than that of the larvae. The adult fly is vegetarian, feeding on decaying fruits and flowers, including the invasive blackberry (*Rubus niveus*) Thunb (Rosales: Rosaceae) [9, 15, 31]. *Philornis downsi* is commonly attracted to a mix of blended papaya and sugar, which is used to trap adult flies (developed by P. Lincango and C. Causton; used by [58], Causton et al. in review). This mix is particularly attractive to adult flies due to the presence

of yeast and fermentation products such as ethanol and acetic acid [59].

suggest that the minimum time for pupation in *P. downsi* is 4–7 days [54].

*DOI: http://dx.doi.org/10.5772/intechopen.88854*

**4.2 Larval development**

however the true cause is unknown.

**5.** *Philornis downsi* **adult behaviour**

*Taxonomic Shifts in* Philornis *Larval Behaviour and Rapid Changes in* Philornis downsi*… DOI: http://dx.doi.org/10.5772/intechopen.88854*

1st instar larva are separated by slightly more than their diameter. First instar larvae lack anterior spiracles (**Figure 1(1B)**). The posterior spiracles of a 2nd instar larva are similarly round with two oval slits; however, the distance between them is two to three times of their diameter (**Figure 1(2D)**; [44]). Anterior spiracles are present during the 2nd instar, and semicircular in shape, lightly pigmented and visible in (**Figure 1(2E)**). 3rd instar posterior spiracular plates are darkly pigmented and round in shape, distinct C-shaped spiracular slits radiate from median ecdysial scar (**Figure 1(3G)**). Pigmentation of the median ecdysial scar is light in early 3rd instar larvae and becoming darkly pigmented later in the stage. Semi-circular anterior spiracles are retained in 3rd instar larvae (**Figure 1(3H)**). Cephaloskeleton morphology differs between instars as outlined in Fessl et al. [44]. Recent studies report a decrease in *P. downsi* puparia size across 2004–2014 [54]. Common et al. (unpublished data), and hence body size is certainly not a useful method to classify instars. In general, it is recommended to use a suite of morphological characters, including anterior and posterior spiracular morphology, to determine the larval instar.

#### **4.2 Larval development**

*Life Cycle and Development of Diptera*

**56**

**Figure 1.**

*cropped and resized in Photoshop CS5.*

on the blood and fluids of their host by penetrating the skin of the nestlings [2, 30]. Larval instar morphological descriptions are given by Fessl et al. [44]. The most distinct character between the instars is the posterior spiracles, which change in colour, shape and number of spiracular slits present throughout larval development [44]. **Figure 1(1A)** shows the posterior spiracles of a 1st instar *P. downsi* larva, characterised by their light pigmentation and two oval slits present [44]. The spiracles of a

*(C) ventral view. (2) second instar: (D) posterior spiracles, (E) lateral view, (F) ventral view. (3) third instar: (G) posterior spiracles, (H) lateral view, (I) ventral view. Obtained by the authors from larvae collected on Floreana Island, Galápagos, Ecuador between 2010 and 2014. The photographs were taken using a visionary digital LK imaging system (dun, Inc) with a canon EOS 5DsR camera and capture one pro 11.3.1, phase one (Flinders University). Images were produced using Zerene stacker 1.04, Zerene systems LLC, software, and* 

*Three larval stages of Philornis downsi. (1) First instar: (A) posterior spiracles, (B) lateral view,* 

The developmental period of *Philornis* larvae is associated with the species' larval feeding habit. For example, time to pupation in coprophagous species takes up to 29 days, but only 4–8 days in subcutaneous species [2, 55]. Larval development periods in free-living species such as *P. downsi* are difficult to determine in the wild as the host nest needs to be dismantled to observe the larvae. Early studies of abandoned or failed nests found 1st instar larvae in nests with 1–3 day old nestlings, 2nd instars in nests with 3–6 day old nestlings and 3rd instars in nests with 3–10 day old nestlings [44]. Larval collections following the cessation of activity at host nests suggest that the minimum time for pupation in *P. downsi* is 4–7 days [54].

Compared with larval development times in the wild, larval development times under laboratory conditions are longer. First attempts to rear *P. downsi* larvae in the absence of a living host had a low success rate, with only three larvae out of 477 reaching the adult stage after a 36 day development time (mean 18 day larval development, 12 day pupation) [56]. As the diet for rearing larvae in captivity was refined, the success rate increased to 10% and larval development time decreased [57]. Development time in the laboratory ranged from 9 to 10 days from larva to pupa [57] with even faster development times occurring as the rearing conditions have improved [pers. comm. P. Lahuatte]. Egg hatch rates in captivity have been high (96%), with most mortality in 1st instar larvae (77%) [57]. Laboratory-based diets that have been developed in the absence of a bird host are primarily based on chicken blood, with more successful diets including hydrolysed protein and vitamin fortification [57]. The lack of keratin in the diet may be causing elevated 1st instar mortality, as 1st instars consume the keratin of the beak in which they reside [44], however the true cause is unknown.

#### **5.** *Philornis downsi* **adult behaviour**

The behaviour of adult *P. downsi* is much less understood than that of the larvae. The adult fly is vegetarian, feeding on decaying fruits and flowers, including the invasive blackberry (*Rubus niveus*) Thunb (Rosales: Rosaceae) [9, 15, 31]. *Philornis downsi* is commonly attracted to a mix of blended papaya and sugar, which is used to trap adult flies (developed by P. Lincango and C. Causton; used by [58], Causton et al. in review). This mix is particularly attractive to adult flies due to the presence of yeast and fermentation products such as ethanol and acetic acid [59].

A one-year study on Floreana Island found that male and female *P. downsi* display dimorphic flight patterns, with females more likely to be caught in high and low traps (2 m, most common at 6–7 m), and males more likely to be caught in traps of intermediate height (4–5 m) [58]. As the pattern of male and female abundance are quadratic opposites, this has tentatively been suggested to be an advantage for females to avoid male flies, as frequent mating in other Dipterans has been found to decrease female reproductive success and lifespan [60, 61]. This flight pattern may also explain why certain host species experience higher parasite intensities, such as the medium tree finch (*Camarhynchus pauper*) Ridgway (Passeriformes: Thraupidae) that has an average nest height of 6 m, thus making it more susceptible to being encountered by female *P. downsi* [58, 62, 63]. However, the factors that cause bird species to experience differing intensities of *P. downsi* are complicated and vary between years. Comparison of flight height in *P. downsi* on different islands is needed to test the generality of this pattern, which may be influenced by average tree height and/or other ecological variables.
