DOI: 10.1111/eea.12325
Feeding location affects demographic performance of
cabbage aphids on winter canola
Ximena Cibils-Stewart1, Brett K. Sandercock2 & Brian P. McCornack1*
1
Department of Entomology, Kansas State University, 123 W. Waters Hall, Manhattan, KS 66506, USA, and 2Division of
Biology, Kansas State University, 116 Ackert Hall, Manhattan, KS 66506, USA
Accepted: 21 April 2015
Key words: finite rate of population change k, life-table response experiment, plant architecture,
population growth, prospective demographic analysis, retrospective demographic analysis,
Brevicoryne brassicae, Hemiptera, Aphididae, Brassica napus, Brassicaceae, stage-structured
population model
Abstract
The cabbage aphid, Brevicoryne brassicae L. (Hemiptera: Aphididae), is a perennial pest that specializes on plants of the Brassicaceae family, attacking winter canola (Brassica napus L.) mainly during
and after flowering. Under field conditions, cabbage aphid colonizes the upper flowering canopy.
Population dynamics of aphids in the flowering canopy could be regulated by differences in either
plant quality (bottom-up) or predatory (top-down) forces. The goal of our study was to determine
the effect of feeding location on cabbage aphid demography. A stage-structured matrix population
model was constructed for aphids restricted to reproductive or vegetative plant tissues of canola. We
found that feeding location had a large impact on demography of cabbage aphid; the finite rate of
increase (k SEM) was higher when aphids were restricted to reproductive tissues, compared to
aphids feeding on vegetative tissues: 1.25 0.01 vs. 1.17 0.01 (leaves). Aphids confined to reproductive tissues with higher k exhibited shorter generation times (T = 14.2 0.2 days) and 53–75%
higher net reproductive rates (R0 = 23.3 1.7) than aphids feeding on vegetative tissues. Prospective analyses showed that there was a nymph-skewed stable stage distribution, and elasticity values
revealed that k is most sensitive to changes in stasis of adults staying in the adult stage and to adult
survival. Retrospective analyses indicated that variation in adult fecundity (value of 0.05) had the
largest effect on population dynamics but collectively, growth of nymphal stage 2–3, 3–4, and 4 to
adult accounted for most of the difference in k between the treatments. Monitoring programs should
target adults and penultimate instars colonizing reproductive tissues of canola plants in the field as
aphids on these plant structures contribute most to population growth.
Introduction
Winter canola, Brassica napus L. (Brassicaceae), is a profitable biofuel crop that has increased in acreage in South
Central USA since the introduction of cold-tolerant varieties. New varieties have allowed growers to rotate canola
with winter wheat, Triticum aestivum L. (Poaceae), the
most abundant crop of this region (Franke et al., 2009;
Ash, 2012). Since its introduction as a new crop, winter
canola has been attacked by a complex of aphid species,
including turnip aphid, Lipaphis erysimi (Kaltenbach),
*Correspondence: Brian P. McCornack, Department of Entomology,
Kansas State University, 123 W. Water Hall, Manhattan, KS 66506,
USA. E-mail: mccornac@ksu.edu
green peach aphid, Myzus persicae (Sulzer), and cabbage
aphid, Brevicoryne brassicae (L.) (all Hemiptera: Aphididae) (Franke et al., 2009; Boyles et al., 2012). When
aphids form dense colonies on developing flowers, yield
losses of up to 70% have been reported if infestations are
left untreated (Boyles et al., 2012). After adoption of seedapplied insecticides for managing early-season turnip
aphids, cabbage aphid has become the most damaging
aphid species colonizing winter canola under mixed
aphid-species infestations (Boyles et al., 2012).
Cabbage aphid is a herbivorous perennial pest restricted
to members of the Brassicaceae and attacks canola mainly
during early flowering and pod development (Boyles et al.,
2012). Cabbage aphid develops through four nymphal instars before reaching physiological maturity and starting
parthenogenetic reproduction (Hughes, 1963). In canola,
© 2015 The Authors. Entomologia Experimentalis et Applicata is published by John Wiley & Sons Ltd on behalf of Netherlands Entomological
Society. Entomologia Experimentalis et Applicata 156: 149–159, 2015
149
This is an open access article under the terms of the Creative Commons Attribution-NonCommercial-NoDerivs License,
which permits use and distribution in any medium, provided the original work is properly cited, the use is non-commercial and
no modifications or adaptations are made.
150 Cibils-Stewart et al.
established populations are mainly composed of apterous
females, but sexual and alatoid individuals are produced
when populations reach high densities. Under asexual and
sexual reproduction scenarios, the fitness of newly deposited apterous nymphs is dependent on deposition sites
selected by their fundatrix mothers on acceptable host
plants. In the field, cabbage aphid is predominantly
observed colonizing the top flowering canopy of the
canola plant (Hopkins et al., 2009). Occurrence of arthropod herbivores in different plant structures is a common
phenomenon in plant-herbivore interactions. Little is
known about the consequence of reproductive cabbage
aphid adults depositing nymphs on suboptimal plant
structures. Reproductive structures of the canola plant
may provide optimal resources for cabbage aphid growth
and development and therefore influence aphid demographic performance (Smallegange et al., 2007; Malik
et al., 2010).
Feeding location has been reported to influence both fitness as well as natural-enemy recruitment for herbivorous
arthropods in other cultivated plant systems including
chili (Idris & Roff, 2002), cotton (Griogolli et al., 2013),
alfalfa (Berberet et al., 2009), tobacco (Athanassiou et al.,
2005; Kavallieratos et al., 2005), sunflowers (Pekar, 2005),
and other brassica crops, including Brassica nigra (L.)
W.D.J. Koch (Smallegange et al., 2007). Conversely, feeding habits of natural enemies can also have disproportionate effects on aphid population growth at locations where
natural enemies are feeding, which has been documented
for soybean aphid in soybean (Costamagna et al., 2013).
In canola, key top-down and bottom-up forces governing
cabbage aphid population dynamics during early flowering and other critical plant developmental stages where
management is most effective (early flowering) are not
well-understood.
The objectives of this study were to use a prospective
demographic analysis and a retrospective life table
response experiment (LTRE) to determine the extent to
which feeding location affects aphid vital rates (survival,
growth, reproduction), and population dynamics. Demographic models based on prospective sensitivity analyses
were used to quantify the potential effects of feeding location on aphid vital rates (reproduction, growth, and survival) and future contributions to population growth rate
or k (Caswell, 1996, 2001). We then used a retrospective
LTRE to decompose the contributions of different vital
rates to variation in k among the experimental treatments.
We hypothesized that reproductive tissues of the canola
plant would support higher population sizes of cabbage
aphid than vegetative tissues. Demographic population
analyses are useful to predict pest outbreaks and dynamics
of colonization in crops (Kasap & Alten, 2006).
Materials and methods
Plant and aphid materials
Canola plants (variety Riley; Kansas Foundation Seed
Service, Manhattan, KS, USA) were seeded in a special
soil mixture that contained all required minerals and
nutrients and maintained in the greenhouse with a L16:
D8 photoperiod for ca. 6 weeks (Johnson-Flanagan &
Spencer, 1994). Soil and photoperiod provided adequate
growth for canola plants to survive the vernalization
process (M Stamm, canola breeder, Kansas State University, pers. comm.). The soil mixture consisted of peat
moss, perlite, gypsum, slow-release fertilizer, and other
micronutrients (proprietary blend, M Stamm). Canola
plants were then artificially vernalized in a growth chamber for ca. 2 months at L12:D12 photoperiod and constant 4 °C to induce reproductive maturity (Murphy &
Scarth, 1994). Vernalized canola plants were watered
daily and used to maintain aphid colonies. Cabbage
aphids used in all experiments were obtained from field
collections but maintained in laboratory-reared colonies
at Kansas State University (KSU) in the Department of
Entomology (Manhattan, KS, USA); aphid species identification was confirmed using alates collected from
founding populations (Blackman & Eastop, 2000). In
April 2011, adult cabbage aphids were collected from
winter canola fields (37°1.35396N, 98°22.01688W and
36°59.79996N, 98°29.03508W) in Barber County, Kansas, transported to the laboratory in coolers, and transferred to vernalized canola plants. Transfer of aphids to
new plants consisted of excising an aphid-infested leaf
and placing it atop of new leaves from vernalized canola
plants to allow independent aphid movement to fresh
plant tissue. Using procedures described by Kos et al.
(2011a), colonies were maintained on vernalized canola
plants at 22 2 °C, 60–70% r.h., and L16:D8 photoperiod. To maintain colony vigor, apterous aphids were
transferred to non-infested, vernalized canola plants
every other week. Voucher specimens (B. brassicae
nymphs and adults) were deposited in the KSU Museum
of Entomological and Prairie Arthropod Research (voucher number 228).
Aphid population dynamics
An exclusion experiment using mesh enclosure cages
was replicated in field and greenhouse trials to assess the
impact of feeding location on aphid population growth
rates in the absence of predation. One trial was setup in
production canola fields at Ashland Bottoms Research
Farm near Manhattan, KS, from 27 April to 18 May
2011, and then repeated under controlled greenhouse
conditions (22 2 °C, 60–70% r.h., and L16:D8
Feeding location affects aphid demography 151
photoperiod) at KSU (Manhattan, KS) from 25 October
to 15 November 2011. Apterous cabbage aphids were
restricted to either the upper flowering canopy or a single leaf in the lower canola canopy using enclosure cages
(23 cm diameter and 71 cm long; Figure 1A) adapted
from Soper et al. (2013). Cages were placed on the same
plant to enable direct comparison of plant location on
aphid population growth, and to control for potential
effects among individual plants in size or nutritional
composition. Each cage consisted of white, no-see-um
mesh (Quest Outfitters, Sarasota, FL, USA) with zippered tops. Zippers provided access to either the flowering raceme or the vegetative leaf after a cage was secured
to a plant structure. The base of each enclosure cage was
secured to the canola plant using 15-cm plastic cable-ties
(Gardner Bender, Butler, WI, USA), which were located
below the last flower of the flowering raceme, or at the
node between the leaf and the plant stem. To allow free
movement of aphids within each cage, we added cylindrical supports made of 14-gauge, galvanized steel wire
(Impex Systems Group, Miami, FL, USA). Supports kept
A
B
C
Figure 1 Pictures depicting enclosure cages deployed to (A)
restrict aphid populations to either reproductive (R) or vegetative
(V) structures, and restrict individual aphids to either (B)
vegetative or (C) reproductive plant tissues.
the mesh from resting on the flowers or the leaves or
disrupting growing aphid populations. In companion
experiments, Cibils-Stewart (2013) showed a lack of cage
effect under field and greenhouse conditions across multiple years using the same cage design. In particular,
cumulative degree-days were the same for aphids
enclosed on reproductive and vegetative plant structures.
A total of 18 (n = 36) and 30 (n = 60) cages per treatment were deployed in the field and greenhouse trials,
respectively.
Populations in each enclosure cage were started with
five newly reproductive, apterous adult cabbage aphids
that were transported to field and greenhouse trials using
2-ml Eppendorf vials (Fisher Scientific, Waltham, MA,
USA). Aphids were transferred to the experimental plants
with fine, camel hair paintbrushes and placed directly on
the canola flowers or leaves. Aphid populations remained
on caged sections of the plant for 3 weeks. Plants within a
trial were arranged in a completely randomized design. At
the end of each trial, the base of each canola plant was
excised and all plant material and attached cages were
placed in a 7.6-l (2 gallons) plastic bag, which was then
immediately stored in a freezer at 20 °C. Freezing
stopped aphid development and nymphal deposition and
allowed for effective counting of cabbage aphid populations at the end of each trial. Plant structures within enclosure cages were removed and all aphids within each cage
were counted in the laboratory using a magnifying glass.
For the greenhouse trial, aphids were categorized into
three morphs based on absence or presence of wing pads
and wings (Blackman & Eastop, 2000). The three morphs
included apterous nymphs and adults, alatoid nymphs,
and alates.
The finite rate of population change was calculated for
the entire 21-day trial (k21) as a ratio of change in aphid
densities at the start (N0) and end (N21) of the field or
greenhouse trials, where k21 = N21/N0. To compare results
between populations and individual aphid experiments,
we then calculated the daily rate of change (kd), where
kd = (N21/N0)1/21. After calculating the finite rate of population growth for each cage location, Student’s paired
t-test (Proc TTEST; SAS 2009; SAS Institute, Cary, NC,
USA) was used to compare aphid populations restricted to
either the top (reproductive) or bottom (vegetative) plant
structures in both trials (greenhouse and field). Lastly,
aphid morphs (apterous nymphs and adults, alatoid
nymphs, and alates) from the greenhouse trial were compared between locations using a Student’s paired t-test
(Proc TTEST; SAS 2009) at a = 0.05. Population structure
of aphids restricted to different plant structures was compared using a Pearson’s v2 test for homogeneity between
feeding locations.
152 Cibils-Stewart et al.
Individual aphid demographics
To model effects of bottom-up forces on individual cabbage aphid, small enclosure cages were used to measure
vital rates (growth, survival, and reproduction) of aphids
feeding on apical reproductive tissues or basal vegetative
tissues of a canola plant. Trials were conducted under
greenhouse conditions (22 2 °C, 60–70% r.h., and L16:
D8 photoperiod) at KSU from 23 September to 28 October 2012. In this experiment, individual aphids were randomly assigned to one of three fixed locations on canola
plants: (1) flowering structure (reproductive tissue), or (2)
top or (3) bottom surfaces of a single leaf (vegetative
tissues) from the mid-canopy of a canola plant. Enclosure
cages (Converters, Huntingdon Valley, PA, USA) followed
the design of Nagaraj et al. (2005) and consisted of a
0.5-cm thick foam rectangle (outside rectangle dimensions: 6.2 9 3.6 cm; inside rectangle dimensions:
5.1 9 2.5 cm) with manufacturer-applied adhesive on
cage tops and bottoms. For cages placed on leaves, preapplied adhesive was then used to secure no-see-um mesh
on one side of the cage. Mesh kept aphids from escaping,
allowed for adequate ventilation, and facilitated repeated
counting of aphid populations. The remaining adhesive
side was secured to the leave on the mid-canopy leaf locations to enclose the aphids (Figure 1B). For aphids
restricted to flowers, two cages were secured together with
a single flower stem between them; no-see-um mesh was
used on the outside of both cages to restrict aphids to
reproductive structures (Figure 1C). Aphids were not
exposed to sticky surfaces to reduce incidental mortality.
Each tissue type was artificially infested with two
apterous adult cabbage aphids (first generation or Gen1)
resulting in 10 cages per treatment (n = 30). On the 1st
day nymphs were produced, ca. 48 h after infestation,
Gen1 adults were removed, and a cohort of five nymphs
(second generation or Gen2) remained in each cage until
nymphs reached the penultimate stage (fourth instar).
Following methods of Chaplin-Kramer et al. (2011), we
kept nymphs together until they reached their penultimate stage to reduce the likelihood of nymphs escaping
individual cages. Only Gen2 nymphs were used to estimate demographic attributes for individual aphids in
this experiment. Nymphal development (instar changes),
adult fecundity, and survival of each Gen2 aphid were
recorded daily for the entire lifespan of each aphid.
Nymphal changes were determined by counting the
aphid exuvia (exoskeleton molts), which were removed
and recorded daily; the number of exuvia was correlated
with nymphal body size to estimate the proportion of
nymphs at a given instar or age. Once Gen2 nymphs
reached the penultimate instar, they were transferred to
individual cages and responses were individually tracked
(n = 50 aphids per treatment location) for the remainder of the experiment (n = 150 total cages). For reproductive Gen2 adults, the number of third generation
(Gen3) nymphs produced by each reproductive adult
was recorded and newly deposited Gen3 nymphs were
removed daily to determine fecundity (total number of
nymphs produced per adult) and reproductive rates
(mean nymphs produced per day per adult).
The life cycle of individual cabbage aphids was categorized into six stage-classes (Figure 2A): four nymphal
stages (collectively pre-reproductive stages) followed by
two adult stages (reproductive and post-reproductive).
The reproductive stage included the number of days that
each female deposited nymphs, whereas the post-reproductive stage included the number of days that adult
females were alive but no longer depositing nymphs. Total
stage duration (number of days) for each nymphal stage as
well as total duration of the pre-reproductive, reproductive, and post-reproductive periods (number of days) were
compared between tissue types using a two-way ANOVA
(Proc MIXED; SAS 2009). In this study, treatment location
(flowers, leaf top, and leaf bottom) was a fixed effect, and
A
B
Figure 2 (A) The life cycle diagram and
(B) corresponding stage-classified
transition matrix for female cabbage
aphids. Stages 1–4 = instars 1–4,
adult = reproductive females, and
post = post-reproductive adult females.
Arrows indicate transitions, where
G = growth (aphid survives from one
stage to the next), S = stasis (aphid
survives and remains in the same stage),
and F5 = fecundity (number of nymphs
per reproductive female at stage 5).
152 Cibils-Stewart et al.
Individual aphid demographics
To model effects of bottom-up forces on individual cabbage aphid, small enclosure cages were used to measure
vital rates (growth, survival, and reproduction) of aphids
feeding on apical reproductive tissues or basal vegetative
tissues of a canola plant. Trials were conducted under
greenhouse conditions (22 2 °C, 60–70% r.h., and L16:
D8 photoperiod) at KSU from 23 September to 28 October 2012. In this experiment, individual aphids were randomly assigned to one of three fixed locations on canola
plants: (1) flowering structure (reproductive tissue), or (2)
top or (3) bottom surfaces of a single leaf (vegetative
tissues) from the mid-canopy of a canola plant. Enclosure
cages (Converters, Huntingdon Valley, PA, USA) followed
the design of Nagaraj et al. (2005) and consisted of a
0.5-cm thick foam rectangle (outside rectangle dimensions: 6.2 9 3.6 cm; inside rectangle dimensions:
5.1 9 2.5 cm) with manufacturer-applied adhesive on
cage tops and bottoms. For cages placed on leaves, preapplied adhesive was then used to secure no-see-um mesh
on one side of the cage. Mesh kept aphids from escaping,
allowed for adequate ventilation, and facilitated repeated
counting of aphid populations. The remaining adhesive
side was secured to the leave on the mid-canopy leaf locations to enclose the aphids (Figure 1B). For aphids
restricted to flowers, two cages were secured together with
a single flower stem between them; no-see-um mesh was
used on the outside of both cages to restrict aphids to
reproductive structures (Figure 1C). Aphids were not
exposed to sticky surfaces to reduce incidental mortality.
Each tissue type was artificially infested with two
apterous adult cabbage aphids (first generation or Gen1)
resulting in 10 cages per treatment (n = 30). On the 1st
day nymphs were produced, ca. 48 h after infestation,
Gen1 adults were removed, and a cohort of five nymphs
(second generation or Gen2) remained in each cage until
nymphs reached the penultimate stage (fourth instar).
Following methods of Chaplin-Kramer et al. (2011), we
kept nymphs together until they reached their penultimate stage to reduce the likelihood of nymphs escaping
individual cages. Only Gen2 nymphs were used to estimate demographic attributes for individual aphids in
this experiment. Nymphal development (instar changes),
adult fecundity, and survival of each Gen2 aphid were
recorded daily for the entire lifespan of each aphid.
Nymphal changes were determined by counting the
aphid exuvia (exoskeleton molts), which were removed
and recorded daily; the number of exuvia was correlated
with nymphal body size to estimate the proportion of
nymphs at a given instar or age. Once Gen2 nymphs
reached the penultimate instar, they were transferred to
individual cages and responses were individually tracked
(n = 50 aphids per treatment location) for the remainder of the experiment (n = 150 total cages). For reproductive Gen2 adults, the number of third generation
(Gen3) nymphs produced by each reproductive adult
was recorded and newly deposited Gen3 nymphs were
removed daily to determine fecundity (total number of
nymphs produced per adult) and reproductive rates
(mean nymphs produced per day per adult).
The life cycle of individual cabbage aphids was categorized into six stage-classes (Figure 2A): four nymphal
stages (collectively pre-reproductive stages) followed by
two adult stages (reproductive and post-reproductive).
The reproductive stage included the number of days that
each female deposited nymphs, whereas the post-reproductive stage included the number of days that adult
females were alive but no longer depositing nymphs. Total
stage duration (number of days) for each nymphal stage as
well as total duration of the pre-reproductive, reproductive, and post-reproductive periods (number of days) were
compared between tissue types using a two-way ANOVA
(Proc MIXED; SAS 2009). In this study, treatment location
(flowers, leaf top, and leaf bottom) was a fixed effect, and
A
B
Figure 2 (A) The life cycle diagram and
(B) corresponding stage-classified
transition matrix for female cabbage
aphids. Stages 1–4 = instars 1–4,
adult = reproductive females, and
post = post-reproductive adult females.
Arrows indicate transitions, where
G = growth (aphid survives from one
stage to the next), S = stasis (aphid
survives and remains in the same stage),
and F5 = fecundity (number of nymphs
per reproductive female at stage 5).
154 Cibils-Stewart et al.
Results
Aphid population dynamics
Daily rate (mean SEM) of population growth (k) for
cabbage aphids was 8% faster for populations restricted to
apical flowers compared to populations restricted to vegetative leaves under greenhouse conditions (1.25 0.01 vs.
1.16 0.01; t = 8.66, d.f. = 59, P<0.001). Daily rate of
population growth (k) for cabbage aphids was
1.17 0.08 for populations restricted to the reproductive
structures and 1.11 0.02 for populations restricted to
vegetative tissue under field conditions (t = 2.41,
d.f. = 35, P<0.05). Mean total numbers of aphids recorded
after 21 days were 1 104 88 and 1 090 227 for populations restricted to the reproductive structures, and
185 26 and 348 104 for populations restricted to
vegetative structures in greenhouse and field trials, respectively. Flower structures supported 5.99 and 3.19 more
aphids than lower leaves of the canola plants after a 3-week
period in greenhouse and field trials, respectively.
Overall population densities were significantly lower
when aphids were restricted to vegetative tissue compared
to reproductive tissues (t = 9.97, d.f. = 59, P<0.001), but
the proportion of apterous nymphs and adults (t = 1.93,
d.f. = 58, P = 0.059) as well as alates (t = 0.46, d.f. = 58,
P = 0.49) within these populations did not differ significantly between feeding locations in the greenhouse trial.
Proportions of the population that were alatoid nymphs,
on the other hand, were significantly higher when aphids
were restricted to reproductive vs. vegetative tissues
(t = 3.84, d.f. = 58, P = 0.0003). Population structure of
cabbage aphid restricted to feeding on the reproductive
parts of the canola plant after a 3-week period comprised
of 72% (786 74) apterous nymphs and adults, 12%
A
(117 14) alatoid nymphs, and 16% (202 34) alates.
Aphids restricted to vegetative tissues of the same plant
had a similar population structure, where 80%
(162 23), 5% (9 2), and 15% (28 5) of all aphids
were apterous nymphs and adults (v2 = 27, d.f. = 59,
P = 1.0), alatoid nymphs (v2 = 13.75, d.f. = 59,
P = 0.55), and alates (v2 = 6.66, d.f. = 59, P = 1.0),
respectively.
Individual aphid demographics
In our trials, 25, 36, and 27 of the 50 aphids survived to
adulthood on the flower, leaf top, and leaf bottom treatments, respectively. Daily nymph counts from surviving
aphids were used for the fecundity analysis. Development
was 20% faster for cabbage aphid restricted to flowers
(10.4 0.8 days) compared to aphids developing on tops
(12.9 0.7 days) or bottoms (12.3 0.8 days) of leaves
(F2,75 = 19.86, P<0.0001). Significant interactions between
plant location and duration of nymphal stage were also
observed (F6,300 = 6.63, P<0.0001) (Figure 3A). Aphids
spent 60–70% less time in pre-reproductive
(8.6 0.1 days) and post-reproductive (6.2 0.5 days)
stages, respectively, compared to reproductive adults
(21.0 0.5 days), regardless of feeding location
(F4,225 = 2.96, P = 0.02) (Figure 3B). Stage duration of
aphids across all plant locations ranged from 7.6 0.1 to
9.2 0.1 days for the pre-reproductive stage, 19.8 0.1
to 21.7 0.1 days for the reproductive stages, and
3.9 0.4 to 8.1 0.8 days for the post-reproductive
stage. Aphids restricted to flowers had significantly shorter
post-reproductive durations than aphids confined to
leaves (F4,225 = 2.96, P = 0.02) (Figure 3B). Hence, mean
( SEM) overall life cycle duration was significantly
shorter for aphids restricted to flowers (31 3 days) than
B
Figure 3 Mean ( SEM) duration (days) of the (A) nymphal stage for cabbage aphids restricted to specific canola tissues (flower, leaf top,
or bottom) and (B) pre-reproductive (Pre-R), reproductive (R), and post-reproductive (Post-R) periods for cabbage aphids restricted to
three canola tissue types (flower, leaf top, or bottom) for a greenhouse trial in 2012. Means within a panel capped with different letters are
significantly different (adjusted Tukey method: P<0.05).
Feeding location affects aphid demography 155
aphids located on upper (39 6 days) or lower leaf surfaces (37 7 days) (F4,225 = 9.98, P<0.001).
Mean daily fecundity values were significantly greater
for aphids restricted to reproductive canola structures
(3.3 0.1 aphids per female per day) compared to aphids
on either top (2.0 0.1) or bottom (2.2 0.1) vegetative
tissues (F2,75 = 19.86, P<0.001). Vital rates of survival, stasis, and mean daily fecundity were pooled across aphids
within the same treatment; and mean values were used for
the prospective analysis (Table 1).
Cabbage aphid restricted to reproductive structures of
the plant had higher demographic performance than
aphids restricted to vegetative plant tissues; population
growth rate (k) was 7% higher for aphids restricted to the
reproductive parts of canola plant (k = 1.25 0.01)
compared
to
aphids
on
vegetative
tissues
(k = 1.17 0.01) (Table 2). Damping ratios (q) and
time of convergence (t20), measured in days, showed that
populations restricted to flowers should converge faster to
the stable-age distribution (5.7 0.1 days) than aphids
restricted to vegetative structures (t20 = 9.4 0.2 and
10.4 0.1 days) (Table 2). Stable age distributions (w)
revealed that first instars are the most abundant stable
stage class among all populations (w>0.30, Figure 4A),
and reproductive values (v, or mean number of offspring
theoretically produced by post-transients) were consistently highest for reproductive adults (v>9), followed by
the fourth instar stage (v 5) (Figure 4B). Life-history
patterns were consistent across treatments. Elasticity
values indicated that k would be most sensitive to changes
Table 1 Vital rates for individual cabbage aphids (n = 50 per
feeding location) feeding on canola, which includes: growth (G),
stasis (S), and fecundity (F, number of females produced by
females per day) rates for nymph (S1–4), adults (S5), and postreproductive (S6) cabbage aphids, restricted to specific tissue
types (flower, leaf top, or bottom) for a greenhouse experiment
in 2012
Vital rate
Flower
Leaf top
Leaf bottom
S1
S2
S3
S4
S5
S6
G1
G2
G3
G4
G5
F1
0.32
0.30
0.41
0.49
0.95
0.74
0.60
0.57
0.41
0.37
0.05
3.25
0.41
0.57
0.43
0.53
0.95
0.88
0.56
0.33
0.39
0.34
0.05
2.04
0.31
0.42
0.53
0.52
0.95
0.85
0.63
0.46
0.29
0.33
0.05
2.15
Table 2 Growth rates and asymptotic matrix properties for individual cabbage aphids restricted to three specific canola plant
structures (flower, leaf top, and bottom) for a greenhouse experiment in 2012. Results obtained from the prospective matrix
analysis are shown, where k = finite rate of population change,
q = damping ratios, t20 = days to model convergence, R0 = net
reproductive rate, and T(d) = generation time in days
Parameter
Feeding location
Mean SEM
k
Flower
Leaf top
Leaf bottom
Flower
Leaf top
Leaf bottom
Flower
Leaf top
Leaf bottom
Flower
Leaf top
Leaf bottom
Flower
Leaf top
Leaf bottom
1.25
1.17
1.17
1.69
1.33
1.38
5.7
10.4
9.4
23.3
15.1
13.3
14.2
16.8
16.4
q
t20
R0
T(d)
0.01
0.01
0.01
0.02
0.02
0.01
0.1
0.1
0.2
1.7
3.2
1.0
0.2
0.8
0.2
95% CI limits
1.23–1.26
1.16–1.19
1.16–1.18
1.65–1.73
1.30–1.37
1.36–1.40
5.5–6.0
9.6–11.4
8.9–9.8
20.2–26.8
10.1–22.2
11.5–15.3
13.9–14.5
15.3–18.5
16.0–16.8
in stasis as adults (s5) and also to survival of adults (a5)
(Figure 5A).
Results from the retrospective analysis comparing the
two leaf treatments and a projection matrix of individual
aphids on reproductive tissue showed that the sum of all
elements in the contribution matrix (∑c) equaled 0.0740;
this sum was a good approximation to the expected treatment effects (Dk: km kr = 0.0739). Positive contributions of improved adult fecundity (F5: 0.049), growth
from nymphal stage 1–2 (g1: 0.007), 2–3 (g2: 0.038), 3–4
(g3: 0.021), and 4 to adult (g4: 0.011), and survival of nymphal stage 3 (a3: 0.005) accounted for nearly all of the difference in k between aphid populations restricted to
canola flowers vs. a leaf on the same plant. These six demographic rates accounted for ca. 74% of the effects of feeding location on fecundity, growth, and survival of cabbage
aphid (Figure 5B). Additionally, there were negative contributions for stasis of early nymphal stages to the variation
in k, where only stasis of the second nymphal stage was
higher for aphids on leaves than flowers (s2 = –0.025)
(Figure 5B).
Discussion
Feeding location on canola had a large effect on the demographic performance of mixed-age and even-aged populations of aphids. Vital rates and the intrinsic rate of growth
(k) were significantly higher for aphids confined to repro-
154 Cibils-Stewart et al.
Results
Aphid population dynamics
Daily rate (mean SEM) of population growth (k) for
cabbage aphids was 8% faster for populations restricted to
apical flowers compared to populations restricted to vegetative leaves under greenhouse conditions (1.25 0.01 vs.
1.16 0.01; t = 8.66, d.f. = 59, P<0.001). Daily rate of
population growth (k) for cabbage aphids was
1.17 0.08 for populations restricted to the reproductive
structures and 1.11 0.02 for populations restricted to
vegetative tissue under field conditions (t = 2.41,
d.f. = 35, P<0.05). Mean total numbers of aphids recorded
after 21 days were 1 104 88 and 1 090 227 for populations restricted to the reproductive structures, and
185 26 and 348 104 for populations restricted to
vegetative structures in greenhouse and field trials, respectively. Flower structures supported 5.99 and 3.19 more
aphids than lower leaves of the canola plants after a 3-week
period in greenhouse and field trials, respectively.
Overall population densities were significantly lower
when aphids were restricted to vegetative tissue compared
to reproductive tissues (t = 9.97, d.f. = 59, P<0.001), but
the proportion of apterous nymphs and adults (t = 1.93,
d.f. = 58, P = 0.059) as well as alates (t = 0.46, d.f. = 58,
P = 0.49) within these populations did not differ significantly between feeding locations in the greenhouse trial.
Proportions of the population that were alatoid nymphs,
on the other hand, were significantly higher when aphids
were restricted to reproductive vs. vegetative tissues
(t = 3.84, d.f. = 58, P = 0.0003). Population structure of
cabbage aphid restricted to feeding on the reproductive
parts of the canola plant after a 3-week period comprised
of 72% (786 74) apterous nymphs and adults, 12%
A
(117 14) alatoid nymphs, and 16% (202 34) alates.
Aphids restricted to vegetative tissues of the same plant
had a similar population structure, where 80%
(162 23), 5% (9 2), and 15% (28 5) of all aphids
were apterous nymphs and adults (v2 = 27, d.f. = 59,
P = 1.0), alatoid nymphs (v2 = 13.75, d.f. = 59,
P = 0.55), and alates (v2 = 6.66, d.f. = 59, P = 1.0),
respectively.
Individual aphid demographics
In our trials, 25, 36, and 27 of the 50 aphids survived to
adulthood on the flower, leaf top, and leaf bottom treatments, respectively. Daily nymph counts from surviving
aphids were used for the fecundity analysis. Development
was 20% faster for cabbage aphid restricted to flowers
(10.4 0.8 days) compared to aphids developing on tops
(12.9 0.7 days) or bottoms (12.3 0.8 days) of leaves
(F2,75 = 19.86, P<0.0001). Significant interactions between
plant location and duration of nymphal stage were also
observed (F6,300 = 6.63, P<0.0001) (Figure 3A). Aphids
spent 60–70% less time in pre-reproductive
(8.6 0.1 days) and post-reproductive (6.2 0.5 days)
stages, respectively, compared to reproductive adults
(21.0 0.5 days), regardless of feeding location
(F4,225 = 2.96, P = 0.02) (Figure 3B). Stage duration of
aphids across all plant locations ranged from 7.6 0.1 to
9.2 0.1 days for the pre-reproductive stage, 19.8 0.1
to 21.7 0.1 days for the reproductive stages, and
3.9 0.4 to 8.1 0.8 days for the post-reproductive
stage. Aphids restricted to flowers had significantly shorter
post-reproductive durations than aphids confined to
leaves (F4,225 = 2.96, P = 0.02) (Figure 3B). Hence, mean
( SEM) overall life cycle duration was significantly
shorter for aphids restricted to flowers (31 3 days) than
B
Figure 3 Mean ( SEM) duration (days) of the (A) nymphal stage for cabbage aphids restricted to specific canola tissues (flower, leaf top,
or bottom) and (B) pre-reproductive (Pre-R), reproductive (R), and post-reproductive (Post-R) periods for cabbage aphids restricted to
three canola tissue types (flower, leaf top, or bottom) for a greenhouse trial in 2012. Means within a panel capped with different letters are
significantly different (adjusted Tukey method: P<0.05).
Feeding location affects aphid demography 157
A
Figure 5 (A) Mean ( SEM) elasticity
values for lower-level vital rates for three
populations of cabbage aphids restricted to
specific canola tissues (flower, leaf top, or
bottom) for a greenhouse trial in 2012, and
(B) contributions of lower-level vital rates
to variation in the finite rate of population
change (k) between cabbage aphid
populations restricted to flowers vs.
vegetative tissues (mean of leaf top and
bottom locations) of canola plants;
positive contributions indicate better
demographic performance in flowers. s,
stasis; g, growth; a, survival; f, fecundity; 14, nymphal stages; 5, adult; 6, postreproductive aphid.
B
graphic parameters that contribute differently to population growth. For instance, a fitness consequence for aphids
feeding in glucosinolate-rich tissues of the plant is reduced
longevity, but mechanisms that effectively sequester glucosinolates may counteract reductions in longevity by shortening generation time and increasing female fecundity
(Agrawal & Kurashige, 2003), resulting in more progeny
produced earlier in the life-history and over a shorter
duration. Shelton (2005) reported that spatial variation in
defenses within or among plant tissues could slow the evolution of resistance to herbivores by creating uneven selection pressure on herbivores and their natural enemies.
Quantification of glucosinolates within different plant tissues and corresponding effects on canola aphids requires
further investigation.
Our prospective analysis showed a nymph-skewed stable stage distribution, which is common in growing populations (Taylor, 1979). Unsurprisingly, reproductive values
were higher for adults and the penultimate nymphal stage
as only adult aphids are reproductive. Elasticity values
showed that k is most sensitive to future changes in stasis
and survival of adults. Even with a nymph-skewed stable
stage distribution, nymphal stages have the highest mortality rates within populations; therefore, adults and penultimate instars are more important in shaping cabbage aphid
population dynamics. Understanding the demographic
composition of aphid populations under field conditions
can guide classical biological control programs. For example, identifying predators or parasitoids that prefer nymphal aphid stages would have the greatest influence on k
(Latham & Mills, 2012). Retrospective analysis indicated
that contributions from nymphal growth stages 2–3, 3–4,
and 4 to adult (Figure 5B) collectively accounted for most
of the variation in k between feeding locations on flowers
vs. leaves; reproductive adults had the single greatest positive contribution to k. We observed that stasis during early
nymph development resulted in negative contributions to
k, which can be explained by higher mortality rates among
early nymphal stages in cabbage aphid. High mortality
(50–72%) during early nymphal stages observed in our
study may be due to host-specific attributes, which has
been documented in other studies. Specifically, Ulusoy &
Olmez-Bayhan (2006) reported host-specific mortalities
ranging from 16 to 88% when aphids were confined to different brassica hosts (broccoli, cabbage, mustard, cauliflower, turnip, and rapeseed); the lowest mortality (16%)
was observed when cabbage aphids were confined to cabbage.
Feeding location influenced demography of aphids
directly, where different plant structures are acting as
either sources or sinks to cabbage aphid. Source-sink relationships within the plant directly affected aphid demography at both the individual and population level. Our result
is a novel finding that helps shape our understanding of
how canola plant structures differentially affect aphid
demographic parameters. To our knowledge, this is the
first study to use a combination of prospective and retrospective demographic analyses to identify vital rates that
contribute most to differences between aphids reared on
different plant structures. Our results indicate that specialist aphids not only have greater growth rates, but also
shorter generation times and higher fecundities when
restricted to reproductive tissues of the canola plant, which
may be a direct outcome of a specialist aphid adapting to
canola secondary compounds. We hypothesize that cabbage aphids developing on flower structures will also contribute more to aphid migration events and colonization
of new canola plants, as the proportion of alatoid nymphs
158 Cibils-Stewart et al.
was significantly higher when aphids were restricted to
reproductive vs. vegetative tissues. Our findings are unsurprising because changes in dispersal polymorphisms are
directly related to aphid population size and to crowding
(Boyles et al., 2012).
Comparative data are limited and it is unknown
whether the same tissue-specific, demographic performance occurs for other aphid species in canola. Study of
canola-aphid interactions has important implications for
pest management programs, especially with potential
changes in population dynamics or species distributions
and development of effective forecasting tools. For example, sampling plans currently used by canola producers
rely solely on whole-plant population estimates. Sampling
plans and corresponding treatment decisions can be
improved by accounting for the unequal aphid growth
rates among feeding locations on the plant. The demographic data presented here in combination with general
field observations can be used to generate more comprehensive population growth models that account for vertical distribution of aphids and can facilitate more timely
management of cabbage aphid in canola.
Acknowledgements
We thank members of the McCornack Field Crops IPM
Lab (KSU) for assistance with field and greenhouse data
collection, and Kris Giles (Oklahoma State University,
OSU) and James Nechols (KSU) for providing feedback
on the experimental design and for reviewing earlier versions of the manuscript. We also appreciate comments
from the anonymous reviewers. We thank Mike Stamm
for providing canola seeds, soil, and growth chamber
space, which were vital the production of vernalized canola
plants and aphid colony vigor. Experiments were a part of
an ongoing collaboration between research scientists at
KSU, OSU, University of Arkansas, and USDA-ARS
(Mericopa, AZ, USA). The project was supported by Agriculture and Food Research Initiative Competitive Grant
no. 011-67009-3014 from the USDA National Institute of
Food and Agriculture and by the Kansas Agricultural
Experiment Station (KAES); this article is contribution 14329-J from the KAES.
References
Agrawal AA & Kurashige NS (2003) A role for isothiocyanates in
plant resistance against the specialist herbivore Pieris rapae.
Journal of Chemical Ecology 29: 1403–1415.
Agrawal AA, Strauss SY & Stout MJ (1999) Costs of induced
responses and tolerance to herbivory in male and female fitness
components of wild radish. Evolution 53: 1093–1104.
Ahuja I, Rohloff J & Bones AM (2009) Defense mechanisms of
Brassicaceae: implications for insect-plant interactions and
potential for integrated pest management. Agronomy for Sustainable Development 30: 311–348.
Ash M (2012) Canola. USDA Economic Research Service. Available at: www.ers.usda.gov/topics/crops/soybeans-oil-crops/
canola.aspx#.UXqqSkoaswc (accessed 10 October 2012).
Tomanovic S
Athanassiou CG, Kavallieratos NG, Tomanovic Z,
& Milutinovic M (2005) Development of a sampling plan for
Myzus persicae (Hemiptera: Aphidoidea) and its predator Macrolophus costalis (Hemiptera: Miridae) on tobacco. European
Journal of Entomology 102: 399–405.
Augustinussen E (1987) The influence of nitrogen fertilization on
growth and development of winter oilseed rape. Danish Journal of Plant Science 91: 301–311.
Berberet RC, Giles KL, Zarrabi AA & Payton ME (2009) Development, reproduction, and within-plant infestation patterns of
Aphis craccivora (Homoptera: Aphididae) on alfalfa. Environmental Entomology 38: 1765–1771.
Blackman RL & Eastop VE (2000) Aphids on the World’s Crops,
An Identification Guide, 2nd edn. John Wiley and Sons,
Chichester, UK.
Boyles M, Bushong J, Sanders H & Stamm M (2012) Great Plains
Canola Production Handbook, pp. 1–60. Kansas State University Agricultural Experiment Station and Cooperative Extension Service, Manhattan, KS, USA.
Caswell H (1996) Analysis of life table response experiments II.
Alternative parameterizations for size- and stage-structured
models. Ecological Modeling 88: 73–82.
Caswell H (2001) Matrix Population Models: Construction,
Analysis and Interpretation, 2nd edn. Sinauer Associates, Sunderland, MA, USA.
Chaplin-Kramer R, Kliebenstein DJ, Chiem A, Morrill E, Mills
NJ & Kremen C (2011) Chemically mediated tritrophic interactions: opposing effects of glucosinolates on specialist herbivores and its predators. Journal of Applied Ecology 48: 880–
887.
Cibils-Stewart X (2013) Influence of Plant Architecture on Tritrophic Interactions between Winter Canola (Brassicae napus),
Aphids (Hemiptera: Aphididae) and Hippodamia convergens
(Coleoptera: Coccinellidae). MSc Thesis, Kansas State University, Manhattan, KS, USA. Available at: hdl.handle.net/2097/
16875 (accessed 18 December 2014).
Costamagna AC, McCornack BP & Ragsdale DW (2013)
Within-plant bottom-up effects mediate non-consumptive
impacts of top-down control of soybean aphids. PLoS one 8:
e56394.
Franke TC, Kelsey KD & Royer TA (2009) Pest Management
Needs Assessment for Oklahoma Canola Producers. EPP7085, Oklahoma Cooperative Extension Service, Stillwater,
OK, USA.
Griogolli JFJ, Souza LA, Fraga DF, Funichello M & Busoli AC
(2013) Within plant distribution of Anthonomus grandis (Coleoptera: Curculionidae) feeding and oviposition
damages in cotton cultivars. Ci^encia e Agrotecnologia 36: 78–
84.
Feeding location affects aphid demography 159
Hopkins RJ, van Dam NM & van Loon JJA (2009) Role of glucosinolates in insect-plant relationships and multitrophic interactions. Annual Review of Entomology 54: 57–83.
Hughes RD (1963) Population dynamics of the cabbage aphid,
Brevicoryne brassicae (L.). Journal of Animal Ecology 32: 393–
424.
Idris AB & Roff MNM (2002) Vertical and temporal distribution
of Aphis gossypii Glover and coccinellid populations on different chilli (Capsicum annum) varieties. Journal of Asia-Pacific
Entomology 5: 185–191.
Johnson-Flanagan AM & Spencer MS (1994) Ethylene production during development of mustard (Brassica iuncea) and
canola (Brassica napus) seed. Plant Physiology 106: 601–606.
Kasap OE & Alten B (2006) Comparative demography of the
sand fly Phlebotomus papatasi at constant temperatures. Journal of Vector Biology 31: 378–385.
Sciarretta A,
Kavallieratos NG, Athanassiou CG, Tomanovic Z,
c V (2005) Seasonal occurrence, distribuTrematerra P & Ziki
tion and sampling indices for Myzus persicae (Hemiptera:
Aphidoidea) and its parasitoids (Hymenoptera: Braconidae:
Aphidiinae) on tobacco. European Journal of Entomology
102: 459–468.
Kos M, Broekgaarden C, Kabouw P, Oude Lenferink K, Poelman
EH et al. (2011a) Relative importance of plant-mediated bottom-up and top-down forces on herbivore abundance on Brassica oleracea. Functional Ecology 25: 1113–1124.
Kos M, Kabouw P, Noordam RA, Hendriks K, Vet LEM et al.
(2011b) Prey-mediated effects of glucosinolates on aphid predators. Ecological Entomology 36: 377–388.
Latham DR & Mills NJ (2012) Host instar preference and functional response of Aphidius transcaspicus, a parasitoid of mealy
aphids (Hyalopterus species). BioControl 57: 603–610.
Malagoli P, Laine P, Rossato L & Ourry A (2005) Dynamics of
nitrogen uptake and mobilization in field-grown winter oilseed rape (Brassica napus) from stem extension to harvest: I.
Global N flows between vegetative and reproductive tissues
in relation to leaf fall and their residual N. Annals of Botany
95: 853–861.
Malik MS, Riley MB, Norsworthy JK & Bridges W (2010) Variation of glucosinolates in wild radish (Raphanusra phanistrum)
accessions. Journal of Agricultural and Food Chemistry 58:
11626–11632.
McCornack BP, Costamagna AC & Ragsdale DW (2008) Withinplant distribution of soybean aphid (Hemiptera: Aphididae)
and development of node-based sample units for estimating
whole-plant densities in soybean. Journal of Economic Entomology 101: 1488–1500.
Mills LS, Doak DF & Wisdom MJ (1999) Reliability of conservation actions based on elasticity analysis of matrix models.
Conservation Biology 13: 815–829.
Murphy LA & Scarth R (1994) Vernalization response in spring
oilseed rape (Brassica napus L.) cultivars. Canadian Journal of
Plant Science 74: 275–277.
Nagaraj N, Reese JC, Tuinstra MR, Smith CM, Amand P et al.
(2005) Molecular mapping of sorghum genes expressing tolerance to damage by greenbug (Homoptera: Aphididae). Journal
of Economic Entomology 98: 595–602.
Pekar S (2005) Horizontal and vertical distribution of spiders
(Araneae) in sunflowers. Journal of Arachnology 33: 197–204.
Pratt C, Pope TW, Powell G & Rossiter JT (2008) Accumulation
of glucosinolates by the cabbage aphid Brevicoryne brassicae a
defense against two coccinellid species. Journal of Chemical
Ecology 34: 323–329.
Ratzka A, Vogel H, Kliebenstein DJ, Michell-Olds T & Kryomann
J (2002) Disarming the mustard oil bomb. Proceedings of the
National Academy of Science of the USA 99: 11223–11228.
Shelton AL (2005) Within-plant variation in glucosinolate concentrations of Raphanus sativus across multiple scales. Journal
of Chemical Ecology 31: 1711–1732.
Slemens DH & Mitchell-Olds T (1996) Glucosinolates and herbivory by specialists (Coleoptera: Chrysomelidae, Lepidoptera:
Plutellidae): consequences of concentration and induced resistance. Environmental Entomology 25: 1344–1353.
Smallegange RC, van Loon JJA, Blatt SE, Harvey JA, Agebirk N &
Dicke M (2007) Flower vs. leaf feeding by Pieris brassicae: Glucosinolate-rich flower tissues are preferred and sustain higher
growth rate. Journal of Chemical Ecology 33: 1831–1844.
Soper AM, Whitworth RJ & McCornack BP (2013) Sorghum seed
maturity affects the weight and feeding duration of immature
corn earworm, Helicoverpa zea and fall armyworm, Spodoptara
frugiperda, in the laboratory. Journal of Insect Science 13: 67.
Stubben C & Milligan B (2007) Estimating and analyzing demographic models using the popbio package in R. Journal of Statistical Software 22: 1–23.
Taylor F (1979) Convergence to the stable age distribution in
populations of insects. American Naturalist 113: 511–530.
Ulusoy MR & Olmez-Bayhan S (2006) Effect of certain Brassica
plants on biology of the cabbage aphid Brevicoryne brassicae
under laboratory conditions. Phytoparasitica 34: 133–138.
Walter AJ & DiFonzo CD (2007) Soil potassium deficiency affects
soybean phloem nitrogen and soybean aphid populations.
Environmental Entomology 36: 26–33.
Winde I & Wittstock U (2011) Insect herbivore counter adaptations to the plant glucosinolate-myrosinase system. Photochemistry 72: 1566–1575.
Winder I, Linton JA, Woolley CC, Perry JN & Holland JM (2012)
The spatial distribution of canopy-resident and groundresident cereal aphids (Sitobion avenae and Metopolophium
dirhodum) in winter wheat. Arthropod-Plant Interactions 7:
21–32.