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Identifier
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Thesis_MES_2022Wi_BoydC
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Title
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IMPORTANCE OF ACCURATE ENVIRONMENTAL CONDITIONS DURING CAPTIVE REARING OF AN ENDANGERED BUTTERFLY (EUPHYDRYAS EDITHA TAYLORI): COLLABORATIVE RESEARCH WITH SUSTAINABILITY IN PRISON PROJECT AND INCARCERATED TECHNICIANS
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Date
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March 2022
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Creator
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Boyd, Carly
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extracted text
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IMPORTANCE OF ACCURATE ENVIRONMENTAL CONDITIONS
DURING CAPTIVE REARING OF AN ENDANGERED BUTTERFLY
(EUPHYDRYAS EDITHA TAYLORI): COLLABORATIVE RESEARCH
WITH SUSTAINABILITY IN PRISON PROJECT AND
INCARCERATED TECHNICIANS
by
Carly M Boyd
A Thesis
Submitted in partial fulfillment
Of the requirements for the degree
Master of Environmental Studies
The Evergreen State College
March 2022
© 2022 by Carly Boyd. All rights reserved
This Thesis for the Master of Environmental Studies Degree
by
Carly M Boyd
has been approved for
The Evergreen State College
by
John C. Withey, Ph.D.
Member of the Faculty
March 18, 2022
Date
ABSTRACT
Importance of Accurate Environmental Conditions During Captive Rearing of an
Endangered Butterfly (Euphydryas editha taylori):
Collaborative Research with Sustainability in Prisons Project & Incarcerated Technicians
Carly M Boyd
Captive rearing is increasingly used as a method to prevent the demise of critically
endangered species. If the conditions under which the captive rearing takes place do not
mimic conditions in the wild, one result may be low productivity and survival of the
species in question. Taylor’s checkerspot butterfly (Euphydryas editha taylori) is an
endangered species endemic to the Willamette Valley-Puget Trough-Georgia Basin
ecoregion of the Pacific Northwest. In 2003, ex-situ conservation programs for E. e.
taylori started at the Oregon Zoo, and expanded to Mission Creek Corrections Center for
Women (Washington) with the Evergreen State College’s Sustainability in Prisons
Project in 2011. The environmental targets for E. e. taylori in captivity were established
based on what is understood to be optimal wild conditions without the extreme that can
occur in the field. To determine the frequency in which the environmental targets were
met, a thorough examination of measured temperature and relative humidity in captivity
was performed. The actual environmental conditions at MCCCW during seven rearing
seasons (2013-2014 to 2018-2019, and 2020-2021) were compared to the environmental
rearing targets to find the percent of time the environmental targets were met and the
percent of days outside of environmental targets for each life stage. Data collected on
productivity and survival—including copulations, oviposition success, egg estimates, and
larval counts at different life stages—were converted to rates and correlated with how
often environmental targets were met in captivity. In addition, since temperature impacts
morphological traits, seasonal morphometric data (adult females’ weight and wing area of
captive and wild butterflies were compared. Environmental targets were infrequently met,
depending on life stages. However, the percentage of time targets were met typically did
not correlate with butterfly productivity, i.e., survival from one life stage to the next. In
addition, captive females weighed more than wild females on average, and had slightly
less wing area. There was also evidence for a slight decreasing trend in wing area, but not
mass, of females over time.
TABLE OF CONTENTS
LIST OF FIGURES ...................................................................................................... VII
LIST OF TABLES ........................................................................................................ XII
ACKNOWLEDGEMENTS ........................................................................................ XIV
ACRONYMS ..................................................................................................................XV
INTRODUCTION............................................................................................................. 1
LITERATURE REVIEW ................................................................................................ 5
LEPIDOPTERA PRESERVATION ............................................................................... 5
Biodiversity Loss and Conservation ........................................................................... 5
Lepidoptera Loss and Conservation ........................................................................... 6
BACKGROUND ON CHECKERSPOTS ...................................................................... 9
Description of Checkerspots ....................................................................................... 9
History of Checkerspot Butterflies ............................................................................ 10
Life History Overview of Checkerspots .................................................................... 12
Environmental Conditions and Checkerspots ........................................................... 14
EUPHYDRYAS EDITHA TAYLORI BACKGROUND ................................................. 19
Euphydryas editha taylori Description & History .................................................... 19
Euphydryas editha taylori Life History..................................................................... 21
Euphydryas editha taylori Habitat Loss & Decline.................................................. 23
Management & Restoration of Euphydryas editha taylori and their Habitat .......... 25
Euphydryas editha taylori Captive Rearing Background ......................................... 28
iv
CAPTIVE REARING ................................................................................................... 31
Captive Rearing & its Risks ...................................................................................... 31
Examples of Captive Rearing Programs................................................................... 34
Environmental Conditions & Captive Rearing ......................................................... 38
METHODS ...................................................................................................................... 43
CAPTIVE REARING PROCESS ................................................................................. 43
Butterfly Technician Environmental Management ................................................... 43
Collecting Wild Adults .............................................................................................. 44
Butterfly Technician Daily Care Procedures ........................................................... 45
CAPTIVE REARING DATA COLLECTION ............................................................. 49
Productivity Outcomes Data Collection & Reporting .............................................. 49
Environmental Data Collection ................................................................................ 50
Morphometric Data Collection ................................................................................. 52
....................................................................................................................................... 52
DATA ANALYSIS ....................................................................................................... 53
Productivity Data ...................................................................................................... 53
Environmental Data .................................................................................................. 54
Spearman’s Rho ........................................................................................................ 56
Morphological Data Analysis ................................................................................... 56
RESULTS ........................................................................................................................ 59
ENVIRONMENTAL DATA ........................................................................................ 59
Overview ................................................................................................................... 59
Temperature .............................................................................................................. 59
v
Relative Humidity...................................................................................................... 60
Outside Temperature Targets ................................................................................... 62
PRODUCTIVITY & SURVIVAL DATA .................................................................... 64
Captive Population ................................................................................................... 64
Wild Population ........................................................................................................ 65
SPEARMAN’S RHO CORRELATION ....................................................................... 67
Captive Population – Productivity/Survival vs. Environmental Targets .................. 67
Wild Population – Productivity/Survival vs. Environmental Targets ....................... 69
MOPHOLOGICAL RESULTS .................................................................................... 71
Weight & Wing Area ................................................................................................. 71
DISCUSSION .................................................................................................................. 73
Adult Butterflies ........................................................................................................ 73
Egg & Larval Stages ................................................................................................. 76
Program Success ....................................................................................................... 77
Recommendations to Environmental Targets & Procedures .................................... 79
CONCLUSION ............................................................................................................... 85
APPENDIX A – ADDITIONAL TABLES ................................................................... 87
APPENDIX B – ADDITIONAL FIGURES ................................................................. 91
REFERENCES.............................................................................................................. 121
vi
LIST OF FIGURES
Figure 1. MCCCW captive rearing greenhouse (left, photo credit: former Butterfly
Program Coordinator) and cold diapause shed (right).................................................... 44
Figure 2. Pictured are the containers in which wild females are transported to MCCCW,
inside a cooler for transport (left) and a wild female feeding (right). .............................. 45
Figure 3. Females inside oviposition chamber, on the mesh closure (left), laying eggs
(middle), and feeding on honey-water solution (right). .................................................... 47
Figure 4. First instar larvae at hatch (left), third instar larvae (middle), and egg and
larval cup set-up (right). ................................................................................................... 47
Figure 5. Diapausing larvae (left) and cold diapause shed set up (right). ...................... 48
Figure 6. 6th instar larvae (left) and pupae (right). Photo credit: Keegan Curry. .......... 49
Figure 7. Wing area measurements (left) and weight measurements (right) for adult
females. Photo credit: former Butterfly Program Coordinator. ....................................... 52
Figure 8. Data points and boxplots of adult female weight (g), by captive (black points)
or wild (grey points) status and across the 2013-14 to 2018-19 seasons (x-axis labels).
The light pink boxes represent 10th and 90th percentiles, the red line the mean, and the
reddish boxes the 95% CI around the mean. .................................................................... 72
Figure 9. Data points and boxplots of adult female wing area (cm2), by captive (black
points) or wild (grey points) status and across the 2013-14 to 2018-19 seasons (x-axis
labels). The light pink boxes represent 10th and 90th percentiles, the red line the mean,
and the reddish boxes the 95% CI around the mean. ....................................................... 72
Figure 10. Example of MCCCW data collection form used during captive rearing. ....... 91
Figure 11. Example snapshot of the hour environmental data analysis into the
percentages of time environmental targets were met. ...................................................... 92
Figure 12. Legend for the percent of time the environmental targets are met. ................ 93
Figure 13. Percent of time environmental targets were met during males for all seasons.
........................................................................................................................................... 93
Figure 14. Percent of time females & oviposition environmental targets were met for all
seasons. ............................................................................................................................. 94
Figure 15. Percent of time eggs & prediapause larvae environmental targets were met
during for all seasons........................................................................................................ 94
vii
Figure 16. Percent of time warm diapause environmental targets were met during all
seasons. ............................................................................................................................. 95
Figure 17. Percent of time cold diapause environmental targets were met for all seasons.
........................................................................................................................................... 95
Figure 18. Percent of time postdiapause environmental targets were met for all seasons.
........................................................................................................................................... 96
Figure 19. Percent of time pupation environmental targets were met for all seasons. .... 96
Figure 20. Legend for the Percent of days the temperature during captive rearing is
outside of the temperature targets. ................................................................................... 97
Figure 21. Percent of days the minimum and maximum daily temperature is outside the
male temperature target. ................................................................................................... 97
Figure 22. Percent of days the minimum and maximum daily temperature is outside the
females & oviposition temperature target. ....................................................................... 98
Figure 23. Percent of days the minimum and maximum daily temperature is outside the
eggs & prediapause larvae temperature target. ............................................................... 98
Figure 24. Percent of days the minimum and maximum daily temperature is outside the
warm diapause temperature target. .................................................................................. 99
Figure 25. Percent of days the maximum daily temperature is outside the cold diapause
temperature target. ........................................................................................................... 99
Figure 26. Percent of days the minimum and maximum daily temperature is outside the
postdiapause temperature target. ................................................................................... 100
Figure 27. Percent of days the minimum and maximum daily temperature is outside the
pupation temperature target. .......................................................................................... 100
Figure 28. Spearman's rho scatterplot matrix for percent males productive versus percent
of time environmental targets were met. ......................................................................... 101
Figure 29. Spearman’s rho scatterplot matrix for percent captive females productive
versus percent of time environmental targets were met.................................................. 101
Figure 30. Spearman’s rho scatterplot matrix for the percent of captive prediapause
larvae versus percent of time environmental targets were met. ..................................... 102
Figure 31. Spearman's rho scatterplot matrix for the percent of captive larvae into
diapause versus percent of time environmental targets were met. ................................. 102
viii
Figure 32. Spearman's rho scatterplot matrix for the percent of captive larvae out of
diapause versus the percent of time environmental targets were met. ........................... 103
Figure 33. Spearman's rho scatterplot matrix for the percent of captive larvae to release
versus percent of time environmental targets were met.................................................. 103
Figure 34. Spearman’s rho scatterplot matrix for the percent of wild females productive
versus percent of time environmental targets were met.................................................. 104
Figure 35. Spearman's rho scatterplot matrix for the percent of the wild prediapause
larvae percent of time the environmental targets were met. ........................................... 104
Figure 36. Spearman's rho scatterplot matrix for the percent of wild larvae into diapause
versus percent of time environmental targets were met.................................................. 105
Figure 37. Spearman's rho scatterplot matrix for the percent of wild larvae out of
diapause versus percent of time environmental targets were met. ................................. 105
Figure 38. Spearman's rho scatterplot matrix for the percent of wild larvae to release
versus the percent of time environmental targets were met. ........................................... 106
Figure 39. Spearman's rho scatterplot matrix for the percent of wild larvae to pupation
(minus the percent of larvae that entered 2nd diapause) versus the percent of time
environmental targets were met. ..................................................................................... 106
Figure 40. Spearman's rho scatterplot matric for the percent of wilds that entered 2nd
diapause (minus the percent of larvae that pupated) versus the percent of time
environmental targets were met. ..................................................................................... 107
Figure 41. Spearman's rho scatterplot matrix for the percent of pupae the successfully
eclosed versus the percent of time environmental targets were met. .............................. 107
Figure 42. Spearman's rho scatterplot matrix of the percent of males productive versus
the percent days outside environmental targets.............................................................. 108
Figure 43. Spearman's rho scatterplot matrix of the percent of captive females productive
versus the percent days outside environmental targets. ................................................. 108
Figure 44. Spearman's rho scatterplot matrix of the percent of captive prediapause
larvae versus the percent days outside environmental targets. ...................................... 109
Figure 45. Spearman's rho scatterplot matrix of the percent captive larvae into diapause
versus the percent days outside environmental targets. ................................................. 109
ix
Figure 46. Spearman's rho scatterplot matrix of the percent of captive larvae out of
diapause versus the percent days above environmental target. ..................................... 110
Figure 47. Spearman's rho scatterplot matrix of the percent of captive larvae to release
versus the percent days outside environmental targets. ................................................. 110
Figure 48. Spearman's rho scatterplot matrix of the percent of wild females productive
versus the percent days outside environmental targets. ................................................. 111
Figure 49. Spearman's rho scatterplot matrix of the percent of wild prediapause larvae
versus the percent days outside environmental targets. ................................................. 111
Figure 50. Spearman's rho scatterplot matrix of the percent of wild larvae into diapause
versus the percent days outside environmental targets. ................................................. 112
Figure 51. Spearman's rho scatterplot matrix of the percent of wild larvae out of
diapause versus the percent days above environmental target. ..................................... 112
Figure 52. Spearman's rho scatterplot matrix of the percent of wild larvae
released/retained versus the percent days outside environmental targets. .................... 113
Figure 53. Spearman's rho scatterplot matrix of the percent of wild larvae pupated versus
the percent days outside environmental targets.............................................................. 113
Figure 54. Spearman's rho scatterplot matrix of the percent of wild larvae return to
diapause versus the percent days outside environmental targets. .................................. 114
Figure 55. Spearman's rho scatterplot matrix of the percent of wild pupae successfully
eclose versus the percent days outside environmental targets. ...................................... 114
Figure 56. Adult female butterfly weights (captive and wild) in the 2013-2014 season
(p=0.03, Wilcoxon rank sum test). The horizontal line inside each diamond is the group
mean. ............................................................................................................................... 115
Figure 57. Adult female butterfly weights (captive and wild) in the 2014-2015 season
(p<0.0001, Wilcoxon rank sum test. The horizontal line inside each diamond is the group
mean. ............................................................................................................................... 115
Figure 58. Adult female butterfly weights (captive and wild) in the 2015-2016 season
(p<0.0001, Wilcoxon rank sum test. The horizontal line inside each diamond is the group
mean. ............................................................................................................................... 116
x
Figure 59. Adult female butterfly weights (captive and wild) in the 2016-2017 season
(p<0.0001, Wilcoxon rank sum test. The horizontal line inside each diamond is the group
mean. ............................................................................................................................... 116
Figure 60. Adult female butterfly weights (captive and wild) in the 2017-2018 season
(p<0.0001, Wilcoxon rank sum test. The horizontal line inside each diamond is the group
mean. ............................................................................................................................... 117
Figure 61. Adult female butterfly weights (captive and wild) in the 2018-2019 season
(p<0.0001, Wilcoxon rank sum test. The horizontal line inside each diamond is the group
mean. ............................................................................................................................... 117
Figure 62. Adult female butterfly wing area (captive and wild) in the 2014-2015 season
(p=0.01, Wilcoxon rank sum test). The horizontal line inside each diamond is the group
mean. ............................................................................................................................... 118
Figure 63. Adult female butterfly wing area (captive and wild) in the 2015-2016 season
(p=0.55, Wilcoxon rank sum test). The horizontal line inside each diamond is the group
mean. ............................................................................................................................... 118
Figure 64. Adult female butterfly wing area (captive and wild) in the 2016-2017 season
(p=0.46, Wilcoxon rank sum test). The horizontal line inside each diamond is the group
mean. ............................................................................................................................... 119
Figure 65. Adult female butterfly wing area (captive and wild) in the 2017-2018 season
(p=0.004, Wilcoxon rank sum test). The horizontal line inside each diamond is the group
mean. ............................................................................................................................... 119
Figure 66. Adult female butterfly wing area (captive and wild) in the 2014-2015 season
(p=0.27, Wilcoxon rank sum test). The horizontal line inside each diamond is the group
mean. ............................................................................................................................... 120
Figure 67. Average temperature over 2021 in Belfair, WA. ........................................... 120
xi
LIST OF TABLES
Table 1. Euphydryas editha taylori captive propagation environmental targets for all life
stages................................................................................................................................. 51
Table 2. Example summary of overall minimum, maximum, and average temperature (ºF)
and relative humidity (%) during captive rearing. This summary is produced for every
life stage in each season. .................................................................................................. 56
Table 3. Overall averages and average ranges of environmental conditions compared to
life stage targets for all life stages in seasons 2013-2014 to 2018-2019 and 2020-2021. 60
Table 4. Average and range of the percent of time the environmental targets are met for
all life stages for seasons 2013-2014 to 2018-2019 and 2020-2021. ............................... 61
Table 5. Average and range of the percent of days outside of the environmental targets
for all life stages for seasons 2013-2014 to 2018-2019 and 2020-2021. ......................... 63
Table 6. Percentage of productivity & survival of captive E. e. taylori populations at
MCCCW for seasons 2013-2014 to 2018-2019. ............................................................... 64
Table 7. Productivity and survival of the wild E. e. taylori populations at MCCCW for
adults to postdiapause stages during seasons 2013-2014 to 2018-2019 and 2020-2021. 66
Table 8. Productivity and survival of the wild E. e taylori populations at MCCCW for
release/retainment to eclosion stages during seasons 2013-2014 to 2018-2019 and 20202021................................................................................................................................... 66
Table 9. Spearman’s rho coefficient for the percent productivity/survival of the captive
population versus percent of time the environmental targets were met for seasons 20132014 to 2018-2019 and 2020-2021. .................................................................................. 67
Table 10. Spearman’s rho coefficient for the percent productivity/survival of the captive
population versus the percent of time outside the temperature targets for seasons 20132014 to 2018-2019 and 2020-2021. .................................................................................. 68
Table 11. Spearman’s rho correlation for the percent productivity/survival of the wild
population versus the percent of time the environmental targets were met for seasons
2013-2014 to 2018-2019 and 2020-2021. ........................................................................ 69
Table 12. Spearman’s rho coefficient between the productivity/survival of the wild
population versus the percent of time outside the temperature target for seasons 20132014 to 2018-2019 and 2020-2021. .................................................................................. 70
xii
Table 13. Median adult female E. e. taylori weights by season at MCCCW, with p-values
from a Wilcoxon rank sum test (see also Appendix B Figures 36-61). ............................. 71
Table 14. Median adult female E. e. taylori wing area by season at MCCCW, with pvalues from a Wilcoxon rank sum test (see also Appendix B Figures 62-66 ...................... ).
........................................................................................................................................... 71
Table 15. Captive rearing productivity and survival targets for seasons 2014-2015 to
2018-2019. ........................................................................................................................ 87
Table 16. Example of a daily minimum, maximum, and average temperature and
humidity table experiences duirng captive rearing. To the right it the comparison of the
absolute targets to the minimum and maximum temperature. .......................................... 87
Table 17. Percent relative humidity averages of the minimum, maximum, and average
percent relative humidity for life stages pupation and males for seasons 2013-2014 to
2018-2019. ........................................................................................................................ 88
Table 18. Percent relative humidity averages of the minimum, maximum, and average for
life stages females and eggs & prediapause larvae for seasons 2013-2014 to 2018-2019
and 2020-2021. ................................................................................................................. 88
Table 19. Percent relative humidity averages of the minimum, maximum, and average for
the life stages warm diapause, cold diapause, and postdiapause. ................................... 88
Table 20. Temperature averages of the minimum, maximum, and average for the life
stages males and pupation. ............................................................................................... 89
Table 21. Overall temperature averages of the minimum, maximum, and average
temperature for life stages females & oviposition and eggs & prediapause. ................... 89
Table 22. Overall temperature averages of the minimum, maximum, and average
temperature for life stages warm diapause, cold diapause, and postdiapause. ............... 89
Table 23. Number of eggs and postdiapause larvae per season. Note, prediapause release
occurred for captive populations, and 20-21 season totals include larvae in the second
greenhouse at MCCCW. ................................................................................................... 90
Table 24. Example of morphometric data summary table. ............................................... 90
xiii
ACKNOWLEDGEMENTS
First, thank you to Dr. John Withey as my thesis reader for your significant assistance
and incredible flexibility over the past year.
I want to acknowledge the years of dedication by Mary Linders, without whom none of
this would be possible. Thank you for your commitment to this work, and I am grateful to
have had the opportunity to learn from you. Thank you to Ronda Naseth with the captive
rearing program at Coffee Creek Correctional Facility in OR, for sharing your expertise
and helping me better understand this work's intricacies. I want to extend my thanks to
others in the teams at the Oregon Zoo, Washington Department of Fish and Wildlife, and
U.S. Fish and Wildlife Service that have devoted years to this work.
I’m exceptionally grateful to have had the chance to work with SPP; it has been an
invaluable experience that I’ll cherish forever. Thank you to Kelli Bush for giving me
this opportunity and being a major source of support for me throughout my time there.
I greatly appreciate the efforts of my fellow Butterfly Program Coordinators who came
before me; thank you for your contributions in molding this program into what it is today.
Thank you to the program liaisons at MCCCW, including the incredibly gracious facility
staff and corrections officers.
I want to recognize the individual Butterfly Technicians Research Specialists—presently
and formerly incarcerated at Mission Creek Corrections Center for Women—who
dedicate their physical, mental, and emotional labor to the recovery of this endangered
butterfly. Without the Technicians' contributions to this program, this thesis would not be
possible. Thank you to my colleagues I worked with directly; I am exceedingly grateful
to have had the opportunity to learn from and collaborate with you.
I would also like to acknowledge the hardships I’ve experienced during my thesis process
and time at MES, and how much I persevered. My experiences here were unexpected,
with physical and mental limitations that inhibited my ability to truly connect with the
incredible communities and individuals I’ve had the pleasure of encountering and
learning from. I’m glad for the opportunities I’ve had for personal growth and to see what
I am capable of by making it through these processes.
I am thankful to Rachel for teaching me the skills necessary to get to this point and
always encouraging me through this process, to Debbie and Bob for treating me like
family, and to Claire and Mairade for sticking with me.
Lastly, I am forever thankful for my husband—whose constant encouragement and
support was imperative to my success—my family, and my cat, all of whom I look
forward to being reunited with after three years of challenging work.
xiv
ACRONYMS
JBLM – Joint Base Lewis-McChord
MCCCW – Mission Creek Corrections Center for Women
PNW – Pacific Northwest
SPP – Sustainability in Prisons Project
USFWS – U.S. Fish and Wildlife Service
WPG – Willamette Valley-Puget Trough-Georgia Basin
WADOC – Washington Department of Corrections
WDFW – Washington Department of Fish and Wildlife
xv
INTRODUCTION
Prairies of the Pacific Northwest (PNW) are home to one of the scarcest endemic
butterflies in one of the rarest ecosystems in North America. Not expected to recover
without human intervention, the endangered Taylor’s checkerspot butterfly (Euphydryas
editha taylori) provides an example of a specialist species diminished beyond the point of
natural reestablishment due to climate change and habitat loss (Stinson, 2005; New,
2014b). Pacific Northwest prairies previously teemed with butterflies in flight,
including E. e. taylori, indicating an abrupt transformation to PNW prairies of the
modern-day (Stinson, 2005; Balke and Fyson, 2014). A prime candidate for captive
rearing and habitat restoration, a single extant South Puget Sound prairie population of E.
e. taylori remains on the Joint Base Lewis McChord artillery range 76 in western
Washington. Due to their diminished status, E. e. taylori receiving listing as endangered
in Washington State in 2006 and under the U.S. Endangered Species Act in 2013
(USFWS, 2013).
Captive rearing for this species was initiated at the Oregon Zoo in 2003, with the
first release occurring on Joint Base Lewis McChord (JBLM) in 2006, continuing
annually thereafter (Grosboll, 2004; Linders, 2007; Linders et al., 2019). Official
husbandry protocols were developed and implemented in 2009, and in 2011 the program
expanded to Mission Creek Correctional Center for Women (MCCCW) through a
partnership with the Sustainability in Prisons Project (Barclay et al., 2009). With much to
uncover about the specific life-history traits and habitat needs of E. e. taylori when
conservation initiated, success prevailed thanks to the vast knowledge established from
studies that focused on other Euphydryas editha subspecies—namely E. e. bayensis in
1
California—and dedicated researchers (Ehrlich and Hanski, 2004b; Grosboll, 2004;
Linders, 2007). While the Oregon Zoo experienced challenging seasons with high levels
of larval mortality during and after the dormant life stage in more recent years, MCCCW
continually sees success which has been partially attributed to captive rearing being
carried out predominantly in greenhouses, providing near-ambient conditions during all
life stages (Lewis et al., 2018). Even with many successful releases, this species still
requires captive rearing to prevent extinction.
Employed to prevent the total loss of critically imperiled species, captive
rearing—when aptly coupled with habitat restoration—provides a recovery opportunity
for at-risk species which may not naturally reestablish in the wild. Invertebrates make
excellent candidates for captive rearing programs, frequently with short life spans, quick
reproductive cycles, and small physical sizes, allowing for smaller captive rearing
facilities and shorter rearing seasons compared to most vertebrates (Hughes and Bennett,
1991; Pearce-Kelly et al., 2007). Concerns around the overall effectiveness of captive
rearing programs arise regarding risks associated with small populations sizes, rearing
conditions diverging from wild conditions, and the potential diseases and illnesses to
spread to the wild population (Snyder et al., 1996; Norberg and Leimar, 2002; Adamski
and Witkowski, 2007). If captive rearing proves successful enough to cease, programs
frequently require continued in-situ conservation intervention to maintain the population
(Pau and Holman, 2019).
Consequences of ineffective captive rearing can include loss of genetic material,
adaptation to the captive environment, inbreeding, decrease in overall survivorship, and
mortality (Snyder et al., 1996; Ballou et al., 2010; Miller et al., 2014). The advantages
2
allotted to invertebrates for successful captive rearing also can allow for rapid
consequences if captive conditions do not adequately mimic wild conditions necessary
for that species to survive and reproduce once released back into the wild (Lewis and
Thomas, 2001; Schultz, Dzurisin and Russell, 2008; Christie et al., 2012). While short
lifespans can be beneficial in producing many individuals for reintroduction from an
endangered invertebrate species, procedures must provide accurate life-history
information and be followed precisely to ensure captive conditions mimic wild
conditions.
Temperature can critically impact the morphological and life-history traits of
butterfly species (Nicholls and Pullin, 2000; Berwaerts, Van Dyck and Aerts, 2002;
Norberg and Leimar, 2002). Maintaining environmental conditions that mimic the natural
conditions a species would experience in the wild is exceedingly crucial for captive
wildlife breeding. For E. e. taylori, captive conditions are maintained based on
environmental targets developed to mimic wild conditions in the prairies while
eliminating extreme conditions that may occur in the wild. Captive rearing conditions get
reported based on environmental captive rearing targets that were officially established
during the 2014 Captive Rearing Meeting—held between Washington Department Fish
and Wildlife, Oregon Zoo, and Sustainability in Prisons Project program partners—and
have been adjusted over the years with current targets found in MCCCW captive rearing
procedures (Lewis et al., 2018; Curry et al., 2020).
No study has yet determined if captive rearing conditions at MCCCW meet these
targets, nor the impacts of the actual environmental conditions on captive rearing
outcomes such as E. e. taylori productivity and survival through all life stages and
3
morphological measurements for adult females. This study will determine how often
these current environmental targets are realized by analyzing the actual conditions during
captive rearing. This information will be used to indicate if correlations exist between E.
e. taylori survival—from copulation and egg-laying through to post-diapause, pupation,
and eclosion—and environmental conditions experienced in captivity. Additionally, the
morphological measurements of the wild adult females—brought to the facility to
produce larvae for captive rearing—and captive-bred adult females will be compared to
support or oppose if selective pressure is acting upon the captive population.
4
LITERATURE REVIEW
LEPIDOPTERA PRESERVATION
Biodiversity Loss and Conservation
Biodiversity loss disrupts internal structures that uphold diverse ecosystems and
contemporary societies, constituting biodiversity preservation an integral and obligatory
part of conservation management plans. Unprecedented rates of biodiversity loss hasten
extinction rates and drove 200 invertebrate species to extinction in one century—
conversely, background extinction rates indicate average loss incurring with one species
lost every 50 years (Ceballos, Ehrlich and Dirzo, 2017). This sixth mass extinction
event—named the Anthropocene extinction due to the overwhelming evidence
designating humanity's pursuit of growth and development primary contributors—
necessitates the commitment to preserve the biodiversity that remains (Agrawal and
Redford, 2009). Worldwide exponential population growth and increased urbanization
coupled with the expanding access to technology and divergence from historically less
invasive resource management tactics in the global north fuels this excessive
consumption and degradation of biodiversity (Wood et al., 2000).
Primary outcomes of this ramped development include land use and
transformation, overexploitation of resources, introducing invasive species, and climate
change, concurrently deteriorating habitat and accelerating loss (Ehrlich and Wilison,
1988; Vijeta, Shikha and Anamika, 2021). Habitat degradation principally drives
biodiversity vulnerability and narrows the prospects for species recovery since underlying
impacts may not immediately present themselves (Tilman et al., 1994; Guardiola et al.,
5
2018). In addition, small or fractured populations have a higher chance of having lower
genetic diversity with no connectivity to other populations' genetic material: a small
population’s gene pool, when cut off from any other source of genetic material, increases
the risks of inbreeding depression and genetic drift which can result in lower fitness for
the populations (Gilbert and Singer, 1973; Frankham, Briscoe and Ballou, 2002; Rochat
et al., 2017).
Modern biodiversity loss prevention methods tend to function retroactively and
the need for intervention generally presents itself once a species reaches a point of critical
concern. The preeminent method for combating extinction is to prevent endangerment
and habitat loss, though, for at-risk species in the present, this alone likely will not lead to
recovery. Habitat degradation often impacts the ecosystem's composition indeterminably
at first, and once species loss accelerates, the impacts of habitat deterioration become
apparent (Kuussaari et al., 2009). This phenomenon, termed an extinction debt,
insidiously delays recovery response time and action, as the impact of the environmental
changes on the residing population do not immediately present themselves yet will
rapidly become apparent as the habitat continues to deteriorate (Tilman et al., 1994;
Kuussaari et al., 2009; Guardiola et al., 2018). Although extinction debts often occur,
New (2014b) discusses how recovery prospects through conservation and habitat
management plans remain possible if the species still endues.
Lepidoptera Loss and Conservation
Lepidoptera—the order of butterflies and moths—hold no immunity to the
unprecedented extinction rates plaquing global biodiversity. In one study of 435 butterfly
species native to Europe, about 83 butterflies (19%) were considered threatened or near6
threatened, 34 species (8%) vulnerable or endangered, and four species (1%) critically
imperiled or extinct (van Swaay et al., 2010). Research conducted at a nature reserve in
Sweden over 50 years revealed that 159 of 597 species (27%) could no longer be found
by 2004, declaring these species extinct (Franzén and Johannesson, 2007). These
examples represent a small proportion of studies that collectively reveal an alarming
downward trend in butterfly diversity worldwide (Sánchez-Bayo and Wyckhuys, 2019).
Efforts to preserve insect biodiversity increased over time, corresponding with an
increase in understanding of the pivotal role many insects play in their ecosystems. The
sheer quantity of insects indicates their importance, suggesting that if insect species
started vanishing at accelerating rates, the impacts would ripple throughout entire
ecosystems (Black, Shepard and Allen, 2001). Ecologists and restorationists recognize
the importance of insects in their ecosystem, however, the average person may not
validate this reality, and minimal public advocacy for insect preservation can lead to
diminutive political and financial support. For example, out of 720 animal species listed
under the U.S. Endangered Species Act, only 13% are insects, even though research
indicates insects make up at least 70% of animal species richness (Ehrlich and Hanski,
2004b; USFWS, 2021).
One of the most popular insect groups, butterflies possess an immense aesthetic
value that contributes to their status of charismatic ambassadors to their ecosystems,
holding the tremendous potential to bring awareness to declining habitats with their
absence (Ehrlich and Hanski, 2004b; New, 2014a). Congruent with this level of attention,
butterflies make up 33% of insect species and 86% of Lepidopteran species, listed under
the Endangered Species Act (T.R. New, 1997; Sharma and Sharma, 2017; USFWS,
7
2021). Some consider butterfly species indicators of their ecosystems' current health
status. Given Lepidoptera species' high vulnerability to deteriorating habitat, research
shows a decline or change in ecosystem operations corresponding to loss of Lepidoptera
species (Erhardt and Thomas, 1991; Cleary, 2004; Sánchez-Bayo and Wyckhuys, 2019).
This preferential treatment allows for faster responses to at-risk butterfly population
declines. At the same time, this promotes further development of insect conservation
studies and model systems for populations through detailed research that can be applied
to other populations outside that species and order (Hanski, Hellman, et al., 2004; New,
2014c).
Industrial agriculture and the resulting pollution from pesticides and herbicides
contribute to Lepidoptera risks, in conjunction with pathogens, host-species loss, invasive
species, and climate change (Pyle, 1976; McLaughlin et al., 2002; New, 2014b). The
combination of Lepidopterans being ectotherms that behaviorally thermoregulate and one
of the most well-studied insect taxa has made many Lepidoptera species indicators of the
impacts that the current climate crisis is having on their habitat and ecosystem. One of the
most prominent phenological responses to climate change involves the asynchrony
between animal development times and host plant development and senescence (Hill et
al., 2021).
Many generalist species are better able to adapt to climate change through
dispersal ability and limited restrictions to specific hosts or nectaring plants. For example,
in response to warming winters, sachems (Atalopedes campetris), a small skipper
butterfly, shift poleward to more suitable habitat amid the climate crisis (Crozier, 2003,
2004). Far from positive, this effect, referred to as ecological drift, leads to generalists
8
replacing specialists, decreasing the biodiversity within the ecosystem (New, 2014b). For
more specialized butterfly species, this level of adaptation to climate change is not
ordinary. Intrinsic barriers often restrict specialists' ability to disperse along with
physiological restrictions regarding suitable habitat and host plant species (Parmesan et
al., 2015; Hill et al., 2021). All-encompassing in-situ and ex-situ conservation methods
used in conjunction make the optimal method for preventing the total demise of these
specialist species.
BACKGROUND ON CHECKERSPOTS
Description of Checkerspots
First described by Boisduval in 1852, 26 different Euphydryas editha subspecies
reside throughout North America (Boisduval, 1852; Pelham, 2012). Named for their
appearance, these graceful butterflies brandish a checkered pattern on their wings that
demands adoration and captivates the imagination (Murphy et al., 2004, p. 18). Medium
in size, checkerspots wingspan average 1.5-3 cm long and primarily have orange, red,
black, or brown colored checkered-pattern (Murphy et al., 2004, p. 22; Pyle and LaBar,
2018, p. 6348). Checkerspots (Euphydryas editha) belong to the Nymphalid family,
commonly referred to as the "brush-foot" butterflies. In adult females, the two front legs
developed over time into sensory organs that assess the acceptability of potential host
plant based on their chemical properties and condition for oviposition (Murphy et al.,
2004, p. 18; Willmott, 2004). Not unique to checkerspot butterflies, it is common to see
checkered patterns in various colors on other species in the Nymphalid family, and the
checkered pattern seems to make prey tracking an arduous task for predators during flight
9
(Nijhout, 1991; Pyle and LaBar, 2018). However, the wings of checkerspot butterflies do
bear a distinct "editha line" that runs between the red or orange bands, distinguishing
them from other similar-looking Nymphalidae species (Murphy et al., 2004, p. 18).
History of Checkerspot Butterflies
About 5 miles from Stanford University campus, on Stanford's Jasper Biological
Preserve, Paul Ehrlich identified a small, checkered butterfly on a hilltop containing a
fragmented patch of grassland encompassed by a border of dense chaparral hillside. Later
named the bay checkerspot butterfly (E. e. bayensis), this subspecies eventually became a
model system for population biology after extensive studies that transpired over
time (Ehrlich, 1961; Agrawal and Redford, 2009). This research exposed the immense
cost resulting from population extinctions; genetic diversity held within distinct
populations grant survival advantages when habitat corridors allow for an exchange of
genetic material between them (Ehrlich and Hanski, 2004a, p. 7). One instance of
utilizing E. e. bayensis research to inform relationships in other species details how the
microclimate impacts dictate phenological relationships between development times of E.
editha larvae and their host plant. This association leads to variation in population
development and survival under a diversity of microclimates and applies to numerous E.
editha ecotypes and other butterfly species that have phenological relationships with their
host species (Hanski, Hellman, et al., 2004). Not only did this species become the
cornerstone for metapopulation research for that time, but the data collected from these
studies have also informed initial life-history traits of checkerspot species across North
America (Ehrlich and Hanski, 2004a; Warren, 2005).
10
Emphasizing the significance of population biology, the sedentary nature of these
butterflies led to disparities in the populations that reside in separate geographical regions
and variation within a single population. Intrinsic barriers to dispersal in checkerspots
limit the flow of genetic material between isolated populations in fragmented habitats.
Studies revealed initial colonies consisting of three populations showed virtually no
migration, and therefore no flow of genetic material, between the populations in question
(Ehrlich, 1961; Singer and Hanski, 2004, p. 184). A more recent study clarified these
findings, showing female Taylor's checkerspots (E. e. taylori) seldom in continuous
flight. Though they found males to be much less sedentary and able to move across
habitat boundaries, they only do so under favorable circumstances and will not disperse
to suitable sites over 100 miles away (Bennett et al., 2013). This inability to maneuver
between fragmented habitats isolates populations further by preventing the establishment
of metapopulations and inhibiting reinstatement after stochastic events (Brückmann,
Krauss and Steffan-Dewenter, 2010; Hanski, 2011).
Selection pressures fluctuate depending on habitat types influencing a
population's genetic makeup, and habitat uniformity varies, leading to genetic phenotypic
divergencies even within a single population (Ford and Ford, 1930; Ehrlich, 1984;
Murphy et al., 2004). At-risk checkerspot ecotypes benefit from their subspecies
classifications, allowing them to receive federally listed under the ESA, however, this
often occurs after populations become critically at-risk. Currently, the federal listing
status for E. e. bayensis is threatened, and E. e. taylori and E. e. quino obtained
endangered listing statuses, all of which benefit from protections that prohibit direct harm
11
and preserve habitats, intending to reestablish metapopulations (Murphy et al., 2004, p.
24; USFWS, 2013, 2021).
Life History Overview of Checkerspots
Recognition and follow-up from the scientific community subsequent to initial
research allowed Euphydryas editha bayensis to become an extensively studied
subspecies; this research constituted most of the early life-history framework for other
checkerspot subspecies (Ehrlich, 1992; Ehrlich and Hanski, 2004b; Pyle and LaBar,
2018). The flight season for checkerspots in lower elevations commences around March
to May, while flight seasons in higher elevations begin around June to August (Ehrlich
and Hanski, 2004b). Mating behaviors vary among subspecies, but males often exhibit
patrolling and perching behaviors to secure a mate, while females remain relatively
sedentary during their search for suitable host plants for oviposition (Bennett, Smith and
Betts, 2012; Bennett et al., 2013). Checkerspots predominantly utilize host plant species
from four families within the subclass Asteridae: Acanthaceae, Asteraceae,
Scrophulariaceae, and Plantaginaceae, with the latter two families containing iridoid
glycosides that, when ingested, will ultimately produce unpalatable individuals as a
predator defense (Murphy et al., 2004, p. 22; van Nouhuys and Hanski, 2004).
Checkerspots use these preferred host plants for ovipositing and larval development,
though larvae host plant preferences tend to be less stringent than adult checkerspots
depending on the subspecies and dispersal abilities (Kuussaari et al., 2004, p. 142).
Ovipositing females lay eggs in masses, on or around host plant leaves. Studies
report E. e. bayensis cluster sizes to vary from 20 to 350 eggs, averaging 40-50 eggs per
cluster, with preliminary research showing that a single female E. e. bayensis can lay up
12
to 1,200 eggs in a lifetime (Labine, 1966, 1968; Singer, 1972; Murphy et al., 2004, p. 25)
Eggs develop over a 13 to 15 day period before hatching into first instar larvae. Larvae of
the same egg clusters create tents of webbing to live collectively and consume host plant
leaves. Larvae continue to feed and develop through instars, the quantity of which varies
depending on the subspecies, with E. e. bayensis larvae known to develop through 3
instars prior to an obligatory diapause. The larvae physically mature and darken in color
over this 3-to-5-week prediapause period, demonstrating their micro-climate
requirements and need for basking behaviors for thermoregulation during adult and larval
life stages (Murphy et al., 2004, p. 22). Larvae also produce setae as they develop and
darken, delicate hair structures all over their bodies that mitigate heat loss (Weiss,
Murphy and White, 1988; Hellmann et al., 2004, p. 47).
Having developed sufficiently before all available host plants senesce—and
maintaining their gregarious lifestyle—checkerspot larvae enter the obligatory dormancy
period in mid-summer or early fall to forgo and survive extreme environmental
conditions (Kuussaari et al., 2004, p. 139). Larvae break diapause—concluded by late
winter rains or melting snow depending on the species and environment—and the
surviving postdiapause larvae continue feeding for a few weeks. Postdiapause larvae can
migrate 10-20 meters per day to find acceptable host plants and foraging conditions to
grow from 3 mg at wake up to the necessary size of 300-500 mg for pupating (Hellmann
et al., 2004, pp. 46–47). If adverse weather conditions arise, checkerspots may reenter
diapause and extend their lifecycle into the next year instead of pupating in the hope of a
higher chance of reproductive success (Kuussaari et al., 2004, p. 139). Postdiapause
13
larvae eventually pupate, often found on vegetation right above the ground, and after a
period eclose as adults to complete their lifecycle (Kuussaari et al., 2004; Potter, 2016).
Environmental Conditions and Checkerspots
Analogous with other insects, checkerspot life-history traits hinge on the
phenological circumstances influencing reproductive success (Taylor, 1981; Hellmann et
al., 2004). Contingent upon synchrony between resource availability and adult butterfly
emergence, the reproductive success of checkerspots depends upon an individual's
opportunities to mate, availability and condition of host plants, ability to circumvent
extreme weather conditions, and offspring survival (Weiss et al., 1993; Kuussaari et al.,
2004). Variation in adult emergence times primarily impacts these phenological factors
and therefore reproductive success; larvae produced by adults that eclose early in the
flight season experience higher survival in most habitat conditions, whereas females that
eclose mid-to-late in the flight season produce larvae with more habitat restrictions and
lower prospects for survival, if at all (Weiss, Murphy and White, 1988; Weiss et al.,
1993; Hellmann et al., 2004, pp. 51–53). In homogeneous habitat settings, emergence
times depend on weather conditions and how rapidly larvae develop, established
primarily by larvae's genetic and phenotypic make-up given the minimal variation in host
plant species availability or microclimate from one host plant to another. However, in
heterogeneous habitat settings, emergence times are more influenced by microclimates
that determine development rates and therefore development timing; in cooler
temperatures, the likelihood for larvae to develop appropriately and in time for winter
diapause decreases, explaining why individuals that enclose later in the season have
lower reproductive success (Hellmann et al., 2004).
14
Emerging early and arduously securing a satisfactory host plant for oviposition
alone does not guarantee larval survival. Two primary causes of larval death arise, one
affecting the species disproportionally more than the other. Predation by spiders, insects,
parasitoids, and vertebrates such as birds are known to feed on checkerspot species,
however, there is little evidence that predation or parasitism contributes to significant
quantities of larval mortality (Boggs and Nieminen, 2004; Kuussaari et al., 2004). The
gregarious lifestyle of checkerspot larvae, the unpalatable nature of larvae if primary host
plants contain iridoid glycosides, and the deterrent of external coloration increase this
species’ likelihood of evading predation (Kuussaari et al., 2004, p. 149). Far more
prevalent in E. editha, larval mortality by starvation critically influences population
dynamics. Preventing starvation in prediapause larvae becomes contingent on meeting
microclimate requirements under which host plants are grown, influencing survival and
development times (Kuussaari et al., 2004, p. 138). For checkerspots, these optimal
microclimate conditions tend to be warm and dry.
The causes and impacts of starvation range depending on the ecosystem and
habitat types, varying by subspecies: in areas where summers are dry and warm, early
host plant senescence and the subsequent pre-diapause larval mortality show less than
10% of offspring making it to adulthood (Kuussaari et al., 2004, p. 149; Singer and
Hanski, 2004). A study conducted by Hellmann (2002) showed high surface temperatures
directly influencing the rate of senescence in Plantago erecta and Castilleja—two host
species utilized by E. editha—though Plantago senesced more rapidly than Castilleja. In
these climates, a conflict arises between larval and host plant requirements, where sunny,
warm weather that accelerates larval development also accelerates host plant senescence.
15
Alternatively, cool and rainy weather that prolongs host plant senescence slows
larval development, delaying diapause and postdiapause. A delicate relationship between
adult emergence times, larvae development, host plant senescence, and environmental
conditions becomes apparent. This correlation principally influences adult emergence
timing for the following season and, therefore, a primary influence on population size
fluctuations in checkerspot species (Singer, 1971, 1972; Kuussaari et al., 2004).
Alternatively, where early senescence infrequently occurs, restricted host plant
availability may lead to larval competition causing overconsumption and eventual prediapause mortality. Larval-host plant relationships often phenological strain, depending
on subspecies and habitat types, increasing in frequency as climate change pushes them
farther out of sync (Singer, 1972).
In addition to temperatures impacting larval and host plant development, other
weather patterns notably influence larval survival by expediting or decelerating larval
maturation in relation to the host species. Early studies of five specific Euphydryas editha
bayensis populations observed distinct waning through all populations following a
drought in CA that spanned from 1975 to 1977 (Ehrlich and White, 1980). While dry and
warm conditions are often optimal for developing many E. editha subspecies, similar to
high temperatures, extreme arid conditions accelerate host plant senescence, leading to
larval starvation (Singer, 1972; Hellmann et al., 2004, pp. 51–53). Desiccation of eggs
and larvae may occur in exceedingly dry conditions, though this is less likely than
prediapause larval starvation and this threshold for desiccation depends on the subspecies
(Kuussaari et al., 2004). In contrast, the research found excessive rainfall spurs
population declines due to considerably increasing development times in prediapause
16
larvae (Dobkin, Olivieri and Ehrlich, 1987). Moisture and precipitation, much like
temperature, create a trade-off between the relatively dry requirements needed for larval
development and the moist conditions that would slow larval development and prolong
host plant senescence.
Habitats with diverse topography result in various microclimates, likewise
prompting severance between larval development and host plant senescence, burdening
postdiapause larvae and pupation development times (Singer, 1972; Weiss, Murphy and
White, 1988; Hellmann et al., 2004). An early study examining the impact of slope
direction on larval survival showed significant differences in temperatures on the south
and north-facing slopes versus flat ground. While north-facing slopes saw ground
conditions analogous to air temperatures, areas of flat ground surpassed air temperature
by 41º-54ºF, and south-facing slopes recording temperatures 68º-86ºF higher than air
temperatures (Singer, 1972; Weiss, Murphy and White, 1988; Hellmann et al., 2004),
showing south-facing slope temperatures transcending flat ground conditions by 27º32ºF. Contrary to initial thoughts, the study predominantly found surviving postdiapause
larvae on cooler north and east-facing slopes. Although the cooler conditions slowed
prediapause larval development, it prolonged host plants' senescence and prevented larval
starvation, allowing larvae to reach diapause eventually. Postdiapause larvae would
emerge on these cooler slopes and exhibit basking behavior, utilizing their black color
and setae to increase body temperatures as much as 50-54ºF above-ground air
temperatures and prevent heat loss (Weiss, Murphy and White, 1988; Hellmann et al.,
2004).
17
In alignment with other life stages, once postdiapause larvae pupate, pupae
development rate corresponds to the microclimate of the slope and the ability efficiently
thermoregulate; the success of pupal development is contingent upon prediapause
dispersal ability and proximity to warmer microclimates (Weiss et al., 1987, 1993; Weiss,
Murphy and White, 1988). Similar to other life stages—specifically more sensitive ones
including adults, eggs, and early instar prediapause larvae—extreme heat can lead to
pupal mortality, with an unpublished study observing wide-spread mortality in pupae on
south-facing slopes during a heatwave where ground temperatures, at times, exceeded
105ºF (Hellmann et al., 2004, p. 47).
All of these environmental factors that influence larval and pupal development
times contribute to the timing of adult emergence, which, as previously discussed,
impacts reproductive success more than any other factor. The advantages allotted to
larvae produced by early-emergence females allows these larvae to circumvent most
other elements that influence survival and population fluctuations (Weiss et al., 1993;
Cushman et al., 1994). This study went on to show that not only does annual average
emergence timing vary by 28 days on the same slope, adult emergence varies nearly 43
days between north and south-facing slopes, with postdiapause larval distribution
impacting the average emergence time for adult butterflies by 10 to 12 days (Weiss et al.,
1993; Hellmann et al., 2004, p. 48).
The severe impacts of extended adverse environment conditions spanning
multiple seasons inevitably lead to severe population declines; prolonged years of
extraordinarily cool and moist or warm and dry conditions destabilize the already delicate
relationship between larval and host plant development so significantly and can bring
18
about regional extinctions (Singer, 1972; Hellmann, 2002; Hellmann et al., 2004, p. 51).
Any outside variable that could impact this dependency may have a similar outcome. For
example, climate change increases variability in weather patterns, and a study looking
into the ramifications of increased inconsistent precipitation patterns on E. e.
bayensis populations indicated a corresponding surge in population size variability that
can quickly induce local extinction (McLaughlin et al., 2002). The primary preventative
method for regional population extinctions involves ensuring heterogeneous habitats have
numerous host species to ensure sufficient host overlap and preventing the degradation of
or reestablishing metapopulations (Singer, 1971, 1972; Fleishman et al., 2000;
McLaughlin et al., 2002; Hellmann et al., 2004).
EUPHYDRYAS EDITHA TAYLORI BACKGROUND
Euphydryas editha taylori Description & History
First described by W. H. Edwards in 1988, Euphydryas editha taylori was named
after a well-known Lepidopterist, George W. Taylor (Guppy and Shepard, 2001). This
subspecies was previously found in prairie habitats from the Willamette Valley in OR
through the Salish Lowlands in WA up to the Georgia Basin in BC—the Willamette
Valley-Puget Trough-Georgia Basin (WPG)—and is one of 15 at-risk species found
throughout this region (Schultz et al., 2011; Pyle and LaBar, 2018). Six of these at-risk
species received federal recognition as candidates for listing, or received listing status as
threatened or endangered under the ESA (Pyle and LaBar, 2018; USFWS, 2021). Not the
only checkerspot species residing in the PNW, the petite E. e. taylori is most similar in
size to the E. e. edithana in comparison to the larger E. e. beani and E. e.
19
colonia subspecies, the latter of which being the largest of the four (Guppy and Shepard,
2001; James and Nunnallee, 2011; Pyle and LaBar, 2018). Possessing modest rounded
wings spanning 2.6-4.3 cm, E. e. taylori also has the darkest coloration of all four PNW
subspecies, with alternating predominantly orange, black, and white bands with a distinct,
primarily orange editha line running through them (Heron, 2011; Potter, 2016).
The only PNW checkerspot subspecies that do not inhabit the cascades, E. e.
taylori's historic range once occurred throughout the WPG Today, most of the remaining
populations reside in Washington. The population of primary focus in this research
occupies Joint Base Lewis McChord artillery range 76 in Pierce County, WA (Stinson,
2005; Potter, 2016). One of only 47 butterfly species previously found throughout this
habitat, this E. e. taylori population dwells within the cool and wet South Puget South
Prairie landscape, an inadequate habitat type for most butterfly species (Dunn and
Fleckenstein, 1997; Pyle and LaBar, 2018). E. e taylori habitat requirements remain
consistent even within a range of habitat types and elevations; suitable habitat
requirements include copious host and nectar plants—preferably from various species—
native grasses, patches of bare terrain, and open structure forbs (Stinson, 2005; Potter,
2016). Defined as prairie-oak habitats, these prairies predominantly contain grasses and
white oak (Quercus garryana). E. e. taylori are not restricted to prairie-oak habitats,
found in glacial outwash prairies, forest balds, oak woodlands, coastal bluffs, and
stabilized dunes (Guppy and Shepard, 2001; Stinson, 2005; Schultz et al., 2011; Potter,
2016; Pyle and LaBar, 2018).
E. e. taylori historically utilized the golden paintbrush (Castilleja levisecta) as a
host plant. Though today they primarily utilize the introduced English plantain (Plantago
20
lanceolata) in addition to harsh paintbrush (Castilleja hispida), slender plantain
(Plantago elongate), sea blush (Plectritic congesta), dwarf owls-clover (Triphysaria
pusilla), blue eye Mary (Collinsia spp.), and owl's clover varieties (Orthocarpus
spp.) (Guppy and Shepard, 2001; Severns and Warren, 2008; Schultz et al., 2011;
Buckingham et al., 2016; Haan, Bowers and Bakker, 2021). An unlikely relationship
arose between the non-native P. lanceolata and endangered E. e. taylori, taking the place
of a primary host species. However, E. e. taylori will utilize native host plants if
available, climate change hastens senescence in many of these native species, causing
misalignment in life-history time-frames between former host species and E. e.
taylori (Buckingham et al., 2016; Haan, Bowers and Bakker, 2021). Primary nectaring
species for E. e. taylori include common camas (Camassia quamash), nineleaf biscuitroot
(Lomatium trieratum), and Puget balsamroot (Balsamorhiza deltoidea) (Stinson, 2005;
Potter, 2016).
Euphydryas editha taylori Life History
Akin to other checkerspot ecotypes, Euphydryas editha taylori has life-history
traits comparable to the well-researched E. e. bayensis: E. e. taylori adults emerge around
mid-April to late-May, and fly until about mid-June, depending on weather and resource
availability (Stinson, 2005; Potter, 2016). As previous studies indicated for E. e.
bayensis, E. e. taylori emergence times also vary greatly depending on weather and
microclimate conditions, host plant availability, geography, and the ever-changing
conditions of climate change that influence larval development times. Females search for
acceptable host plants for oviposition during this flight period—predominantly
utilizing P. lanceolata—while males perch and patrol for females to mate with (Murphy
21
et al., 2004; Bennett, Smith and Betts, 2012). Once females have selected a host plant for
oviposition, they will lay eggs in clusters on or around host plant leaves. E. e.
taylori females usually lay multiple egg clusters over the flight season and conclude their
life soon after, completing the species' one-year life cycle, making them
univoltine (Potter, 2016). Eggs develop over 8-14 days before hatching around mid-June
(Barclay et al., 2009; James and Nunnallee, 2011; Curry et al., 2020).
Gregarious larvae feed on host plants and disperse minimal distances, as needed,
in search of more food or optimal conditions for development. They develop from first
instar at hatch through to fifth instar, slowing eating, molting, and developing new skins
at the onset of each instar (Barclay et al., 2009; Potter, 2016). Once larvae reach fifth
instar—about a month after hatching, around July-August—larvae will soon stop eating
entirely and enter diapause, a dormant phase that, depending on development times,
aligns with host plant senescence to prevent larval starvation and allow them to survive
through the winter (Hellmann et al., 2004; Kuussaari et al., 2004). Around late February
to early March, environmental conditions trigger larvae to break diapause and resume
feeding for development as postdiapause larvae. Postdiapause larvae have a higher
capacity to disperse in search of proper host plant and microclimate conditions if needed,
and about late-March to early-May, larvae will pupate (Hellmann et al., 2004; Stinson,
2005; Potter, 2016). Pupation development times can vary greatly depending on
environmental conditions, and adults will eclose anywhere from two to six weeks after
pupation, emerging for the flight season (Barclay et al., 2009; Potter, 2016).
22
Euphydryas editha taylori Habitat Loss & Decline
Conservationists witnessed drastic declines in Euphydryas editha
taylori populations following the dramatic loss of grasslands state and nationwide due to
this species' immense sensitivities to habitat changes and disturbances. In Washington
state alone, prairies once covered over 150,000 acres of land and a meager 8% of
grasslands remaining today, with most composed of only 25% native grasses (Stinson,
2005; Peter and Harrington, 2014). Nationally, the primary reason for grassland loss
occurs through land transformation for other uses, primarily urban spread and conversion
to farmland and pastures; known for their productive soils, prairies require little effort to
convert for agricultural purposes since they do not require clearcutting, allowing the
minimal barrier to entry for development (Dunwiddie and Bakker, 2011). Urban
development also contributes to the loss of prairie-oak habitats and grasslands. The
dominant habitat type throughout the Willamette Valley-Puget Trough-Georgia Basin
consists of prairies and grasslands. Though the WPG accounts for less than 4% of land in
the PNW region, 75% of the population in this region occupy counties or districts that
entirely or partially overlap with the WPG (Dunwiddie and Bakker, 2011).
Wildfire suppression accounts for much of the remaining prairie deterioration,
prior to colonization throughout North America, many First Nations managed grasslands
within the Coast Salish region of what is now known as the northern portion of the PNW
in the United States and BC in Canada (Wonders, 2008). One of the most notable First
Nations management practices included regular controlled burns of grasslands which
allowed game management, hunting, traveling, and cultivations of plants for food,
medicine, and basketry, contributing significantly to maintaining cultures and economies
23
(Agee, 1989, 1996; Boyd, 1999; Hamman et al., 2011; Ryan, Knapp and Varner, 2013).
Following colonization that brought disease, war, genocide, and the imposition of EuroAmerican culture on all remaining native peoples, controlled burns ceased in the 1800s,
and the succession and eventual occupation by Douglas-firs that followed turned many
grasslands into forests (Hamman et al., 2011; Peter and Harrington, 2014).
In recent years, western cultures gained a better understanding of prescribed burns
as a restoration tool for managing ecosystems by looking to these historical methods.
Western restorationists worked with and learned from First Nations people to reintroduce
these management tools under the designation "Traditional Ecological Knowledge" to
reverse some prairie succession and degradation (Hamman et al., 2011; Trager and
Daniels, 2011). Other causes of prairie loss nationwide include habitat fragmentation,
loss of native fauna, the introduction of non-native and invasive fauna, resource
extraction, and military training (Stinson, 2005).
Historically, 45 E. e. taylori population sites existed in WA, and 25 other
population sites existed between OR and BC. In response to habitat loss and
fragmentation, E. e. taylori populations began to decline dramatically by 1991, while just
11 years prior, reports describe dense clusters of E. e. taylori found in OR, providing an
example of an extinction debt evoked by habitat degradation beginning to catch up to the
subspecies by the early 1990s (Dornfeld, 1980; Stinson, 2005). Populations in WA and
BC saw analogous declines in E. e. taylori populations, and information available today
shows only eight extant populations persist in WA (Holtrop, 2010; Holtrop, Hays and
Potter, 2013; Potter, 2016), three populations in OR (Warren, 2005; Page et al., 2008)
and one population in the Denman Island of BC (Guppy and Shepard, 2001; Balke and
24
Fyson, 2014). In WA, most extant E. e. taylori populations dwell around the forests of the
northeastern Olympic Peninsula, utilizing habitat and host plants that grow in forest
clearings and forest balds (Potter, 2016). These forest populations comprise 6 of the eight
total WA populations with one additional population off the Sequim coast, north of the
forest populations, utilizing dunes for habitat. The remaining singular population inhabits
the prairies of JBLM, approximately 110 miles away from the closest extant populations
(Holtrop, Hays and Potter, 2013; Potter, 2016; Linders et al., 2019). A stark contrast to
the 32 historic populations throughout the South Puget Sound prairies, this isolated
population would perish without intensive intervention (Stinson, 2005; Holtrop, 2010).
This dire situation culminated in the listing of E. e taylori, not only as an endangered
species under the Endangered Species Act in 2013 but prior to that, E. e. taylori acquired
endangered listing status in Canada by 2011 and WA by 2006 (Heron, 2011; USFWS,
2012; Potter, 2016). Considering this species significantly reduced population size and
unfeasibility to repopulate naturally, the best option to prevent the total demise of this
delicate butterfly is through all-encompassing in-situ and ex-situ conservation methods.
Management & Restoration of Euphydryas editha taylori and their Habitat
Once the threat to Euphydryas editha taylori's persistence was deemed severe
enough for state and federal agencies to enable listing status, recovery plans for E. e.
taylori and their habitat began development. A massive undertaking, numerous
organizations and agencies collaborate to form partnerships, including the Cascadia
Prairie Oak Partnership, which works throughout western WA and OR to restore and
preserve WPG prairies (Dunwiddie and Bakker, 2011). This cooperative includes state
and federal agencies, universities, municipalities, non-profit organizations, and private
25
landowners, including First Nations peoples, USFWS, WDFW, Center for Natural Lands
Management, Department for Natural Lands Management, The Nature Conservancy, and
Sustainability in Prisons Project (Hamman, 2018). Primary tenets of prairie restoration
focus on preventing succession and infringement of invasive species through mowing,
prescribed burns, and the practice of flagging and removing or spraying invasive and
non-native species with herbicides (Hamman et al., 2011; Schultz et al., 2011).
Balancing habitat restoration efforts and endangered species recovery proves
arduous, especially in the case of sensitive or specialized native plant and animal species.
Considered one of E. e. taylori's historic host species, E. e. taylori will utilize
available C. levisecta for oviposition and larval development, however, E. e.
taylori shows lower survival in early instars when utilizing this species (Buckingham et
al., 2016; Haan, 2017). Some speculate whether a loss of adaptation occurred, limiting E.
e. taylori's ability to utilize C. levisecta effectively and restricting habitat management
efforts. However, earlier studies show little evidence supporting this other than limited
interactions in the wild between the two species, which suggests these two species
possibly are not historically linked to the extent previously described (Haan, 2017). A
more recent study showed lower oviposition preference for C. levisecta, indicating a
change from previous studies, with larval growth limited on Castilleja when they
consumed predominantly leaves instead of bracts (Haan, Bowers and Bakker, 2021).
Presently, captive and wild E. e. taylori primarily utilize P. lanceolata for oviposition and
larval development, creating contention between E. e. taylori recovery that relies on this
non-native species and prairie recovery that revolves around returning South Puget Sound
prairies to native plant communities (Severns and Warren, 2008; Dunwiddie and Rogers,
26
2017). For this reason, efforts to restore prairies are strategic, careful, and never entirely
based around the threatened species in question also considering native species in other
portions of the ecosystem.
Essential components of prairie habitat requirements exist to ensure E. e.
taylori recovery, and specific habitat management plans were established and
implemented to benefit the endangered butterfly and the prairie ecosystem. These include
large, open areas that are well-connected to itself and other suitable adjacent sites, bare
ground with surrounding herbaceous vegetation, high-density patches of acceptable host
and nectar plant, and some topographic variety to allow for minor microclimate
differences (Stinson, 2005; Severns and Grosboll, 2011; Potter, 2016). Other
management efforts revolve around propagating and planting native grasses and forbs
and non-native English plantain and removing invasive species such as Scotch broom
(Cytisus scoparius) and Himalayan blackberry (Rubus armeniacus) (Potter,
2016). Prairies open-areas make them especially at risk for invasive species to move in,
such as false brome (Brachypodium sylvaticum) reported on JBLM lands for the first time
in 2021 (Hamilton-Wissmer, 2021).
Weather events and climatic change alter the relative timing of E. e. taylori and
host plant development, causing them to be out of sync, which shows a need to establish
multiple host species in South Puget Sound prairies to serve as a buffer for early, mid,
and late E. e. taylori emerging into flight season (Singer, 1972; Schultz et al., 2011).
Previously, little was known about the distribution and abundance of E. e. taylori or their
habitat requirements, but in-depth surveying and population monitoring began in the late
1990s and continues today (Morgenweck and Dunn, 2003; Stinson, 2005). There is a
27
better understanding of E. e. taylori's current status and habitat needs, which represents
the best idea of what restoration to a natural landscape should look like in this ecosystem.
However, there is still no definite characterization of what ideal habitat restoration should
look like for this species, this habitat, and other species present in it.
Euphydryas editha taylori Captive Rearing Background
Not expected to repopulate historical occupation sites naturally due to how
fragmented and diminished Euphydryas editha taylori populations became, captive
breeding and captive rearing began work in conjunction with habitat restoration. Exsitu methods supplement the wild E. e. taylori population to prevent complete extinction.
In 2004, WDFW and the Oregon Zoo partnered to develop initial captive rearing and
translocation methods for E. e. taylori, eventually finding that post-diapause larvae are
the most viable life stage for translocation and reintroduction (Grosboll, 2004; Linders,
2007; Potter, 2016). Since only a single naturally-occurring E. e. taylori population
remains in the South Sound prairies, original trials and following efforts rely entirely on
founding individuals from the population on JBLM artillery range 76 (Potter, 2016;
Linders et al., 2019).
These initial captive rearing trials showed success and informed most of the
original captive rearing protocols, demonstrating an increase in survival and production
(Grosboll, 2004). The first reintroduction occurred on JBLM in 2006, occurring annually
thereafter (Linders et al., 2019). These reintroductions occur on four sites within South
Puget Sound prairies—two located on JBLM and two in Thurston County—in the hopes
of eventually developing into a metapopulation. Eventually, established methods for
breeding and rearing E. e. taylori were incorporated by the Oregon Zoo conservation
28
research team working with WDFW under the approval of USFWS (Barclay et al., 2009).
Captive rearing at the Zoo continued from its initial start in 2003 until 2020, when the
program ceased due to funding, impacts of COVID-19, and the seeming improvements of
wild E. e. taylori populations. In 2011, the Sustainability in Prisons Project—a
partnership between The Evergreen State College and WA Department of Corrections—
collaborated with WDFW, the Zoo, and USFWS to begin a captive rearing program
for E. e. taylori in Mission Creek Corrections Center for Women.
Bringing incarcerated individuals closer to nature in it of itself is a net positive.
SPP provides education opportunities, job training, and networking connections—all in
relation to sustainability and the environment—to the carceral system in WA. SPP’s
contributions carry such influence, other states followed suit throughout the country
(Kaye et al., 2015). SPP awards certifications to incarcerated individuals who meet
specific education and experience criteria for some programs, which designates them 15
college credits at The Evergreen State College upon enrollment after release from prison.
SPP and the co-director, Kelli Bush, strive to expand education opportunities and
currently, the SPP team of managers and master's student employees are working to
establish a consistent education curriculum that will allow incarcerated program partners
to earn college credits from TESC while incarcerated (pers. communication, Kelli Bush).
Research carried out by TESC masters students investigating the impacts of SPP
programming on current and formerly incarcerated individuals continues to grow, and
findings indicate positive participant experiences (Clarke, 2011; Weber, 2012; Gallagher,
2013; Webb, 2016; Gilliom, 2017; Passarelli, 2017). However, Webb (2016) expressed
skepticism of the overall influence by discussing the continued existence of the Prison
29
Industrial Complex: a system described by Alexander (2010) as derived from a history of
Jim Crow policy and slavery, and one that disproportionately impacts black people and
all people of color and allows various institutions to benefit from mass incarceration
(Alexander, 2010; FBP, 2022). This context leads some to believe efforts to achieve
sustainability more closely resemble greenwashing, for example Jewkes and Moran
(2015) state that reducing prison population sizes would have a more substantial
environmental and social impact while green prisons become more productive and
competitively efficient. Given the historical and ethical background and the variety of
opinions on the carceral system, the impacts programs have on the individual participants
cannot go overlooked. In addition to growing reports of positive experiences, the job
training, and education opportunities, SPP contributes to post-release support systems, all
of which has been found improve employment opportunities upon release and to reduce
recidivism (Kaye et al., 2015).
At MCCCW in 2011, the first captive rearing season took place in a 3 x 7.3 m
glass greenhouse, and the program saw success comparable to the Zoo (Linders and
Lewis, 2013). Incarcerate butterfly technicians—usually trained by Oregon Zoo keepers
or graduate students employed as Butterfly Program Coordinator with Sustainability in
Prisons Project—carry out daily care of the endangered E. e. taylori. The MCCCW
captive rearing program has expanded over the years; currently carried out in two large
greenhouses with the goal of producing 5,000 postdiapause larvae per greenhouse,
though the program has not yet reached this capacity due to the continuing impacts of
COVID-19. Population supplementation has prevented the total demise of this species
thus far; however, the long-term fate of E. e. taylori is not yet secure. In order for
30
reintroduction sites to be considered reestablished, during the flight season, surveyors
must see over 250 adults in one day over 50 acres of space for five consecutive years,
happening in conjunction with five years of successful releases, followed by five years of
surveilling the prosperous population (Stinson, 2005; Potter, 2016).
CAPTIVE REARING
Captive Rearing & its Risks
Some at-risk populations regress beyond the point of natural recovery, requiring
off-site management. Conservationists typically only resort to captive rearing when a
species becomes critically at-risk in attempt to prevent unnecessary extinction
(Engelmann and Engels, 2002). Ex-situ conservation—which involves rearing individuals
in captive conditions that mimic wild conditions—became common practice over the
years to mitigate the unprecedented biodiversity loss occurring worldwide. In this
practice, the captive stock is either reared or bred to produce offspring with the goal of
eventual release during the life stage with the highest chance to survive such
reintroduction back into that species’ wild habitat (Engelmann and Engels, 2002).
However, not a catch-all solution for wide-spread species endangerment, the
implementation of captive rearing remains reserved for dire circumstances of critical
endangerment. It provides a short-term solution to prevent the total eradication of at-risk
species due to dramatic declines caused by habitat fragmentation and
degradation (Hughes and Bennett, 1991; Caughley and Gunn, 1996; Crone, Pickering and
Schultz, 2007).
31
Reintroductions fail to find success when restorationists fall short when
attempting to address habitat degradation concerns alongside implementing exsitu conservation methods; captive rearing efforts have the highest chance for success
when coupled with habitat restoration and management plans (Morton, 1983; Adamski
and Witkowski, 2007). In-situ and ex-situ conservation methods used congruently allow
restorationists to invest in and ascertain valuable life history information of other species
throughout the habitat, leading to more inclusive management plans (Ehrlich and Hanski,
2004b; Grosboll, 2004; Daniels et al., 2020). Since primary resources initially tend to go
towards charismatic megafauna, this allows for the chance to increase education and
outreach into the surrounding community that helps support public engagement in
conservation efforts to benefit more than the species in question.
Risks heighten when implementing ex-situ conservation methods with
significantly diminished endangered species populations, and these methods alone cannot
assume to completely fix these dire situations once a population’s genetics become
severely constrained (Rahbek, 1993; Sigaard et al., 2008). Primary constraints of this
practice that could lead to increased risk include financial and physical resources
restrictions, the possibility of spreading diseases and parasites from captive to wild
populations, and genetic hazards derived from small population sizes. These practices
frequently initiate with a diminished founder population and, therefore, a small gene pool
given the nature of critically endangered species (Ballou et al., 2010). Usually,
constraints limit the understanding of the diminished founder population’s genetic
makeup, impeding the ability to track lineages and relatedness to prevent inbreeding
(Hedrick and Hurt, 2012; Miller et al., 2014). In order to feasibly circumvent inbreeding
32
with minimal genetic information, best practices comprise translocation from larger,
more established populations to be reintroduced into smaller, less stable populations
(Crone, Pickering and Schultz, 2007). However, minor or significant genetic differences
often arise between segregated populations depending on habitat types that could affect
fitness if translocated to other habitat types (Frankham, Briscoe and Ballou, 2002; Crone,
Pickering and Schultz, 2007).
Traits that allow successful survival and reproduction in the wild can be
unintentionally subject to selection during captivity (Nylin and Gotthard, 1998; Norberg
and Leimar, 2002; Frankham, 2008). Once a critically endangered population is deemed
suitable for captive rearing and eventual reintroduction, captive conditions must mimic
wild conditions sufficiently to prevent selective pressure from causing adaptations to
captive conditions (Frankham, 2008). Selection pressure can lead to naïve individuals,
less adept at surviving upon release (Sutherland, 1998; Bryant and Reed, 1999; Stockwell
and Weeks, 1999; Frankham, 2008). These genetic adaptations can occur exceedingly
rapidly, observed to occur in as little as one generation (Dzurisin, 2005; Crone, Pickering
and Schultz, 2007; Christie et al., 2012). Due to financial and resource constraints, it is
rare to assess and compare captive and wild individuals of a species to understand if
adaptations to captivity are occurring and how that impacts survival (Schultz, Dzurisin
and Russell, 2008). The risks of working with a small gene pool are compounded and can
lead to further loss of genetic materials in the captively reared population, including an
increase in genetic homogeneity and loss of rare alleles (Snyder et al., 1996; Crone,
Pickering and Schultz, 2007; Schultz, Dzurisin and Russell, 2008).
33
Unfortunately, multiple captive rearing and captive breeding programs show
decreased fitness within the reintroduced individuals when post-release monitoring
occurs (Snyder et al., 1996; Christie et al., 2012). In order to maintain the genetic
integrity of the captively reared population and prevent the production of less-fit
individuals, captive conditions must mimic wild conditions as closely as possible,
including seasonal habitat environmental conditions (Frankham, 2008). Challenges arise
when working to understand what exact environmental requirements entail, in addition to
other factors—habitat size, conditions, connectivity, and weather conditions that
determine the success of that captive rearing reintroduction—and it becomes difficult to
understand how minor misalignments could impact the likelihood of reintroduction
success after captive rearing (Oates and Warren, 1990; Hanski, Ehrlich, et al., 2004). The
extent of this trial-and-error period prolongs the length of the captive rearing program,
which can lead to long-term supplementation of captive individuals into wild populations
as factors of captive rearing and habitat management get refined. These long-term
programs go against long-standing recommendations that these practices are interim
programs to prevent the total demise of the population and only be used as a last resort
(Pyle, 1976; Hughes and Bennett, 1991).
Examples of Captive Rearing Programs
Even with the inherent risks, captive rearing holds considerable potential to save
invertebrates specifically from extinction for a plethora of reasons. Invertebrates tend to
have diminutive sizes, brief life spans, and rapid reproduction times that allow captive
rearing programs to be more financially attainable and swiftly produce wildlife for
release (Hughes and Bennett, 1991; Pearce-Kelly et al., 2007; Linders and Lewis, 2013)
34
However, genetic risks quickly compound in invertebrates due to these life history traits,
allowing for rapid onset of genetic divergences that could affect the captive population.
Even with the risks, the benefits of preventing the total demise of endangered species
outweigh potential hazards; butterfly captive rearing and breeding programs have
increased exponentially since the early 2000s, with the expansion occurring at a faster
rate in the U.K. compared to the U.S. (Schultz, Russell and Wynn, 2008). Exsitu conservation is recommended for about 50% of threatened or endangered butterfly
species under the ESA, with half focusing on captive rearing and the other half on captive
breeding (Schultz, Russell and Wynn, 2008).
A remarkably high-profile species, monarch butterflies, Danaus plexippus, inspire
and enlighten the public given their beauty and fascinating life history. Though
infrequent, a charismatic species occasionally benefits from the public widely supporting
its conservation. Often considered flagship species, these designations and public
awareness help restoration progress and research by accumulating financial aid and
resource support towards the species recovery (Tim R. New, 1997; Barua et al., 2012;
New, 2014c). Public concern also leads to outreach, including volunteers participating in
the species recovery process. Many citizens began rearing or purchasing monarch
butterflies from commercial breeders to eventually release them in the wild when this
species became unstable and at-risk in some areas (Davis, Smith and Ballew, 2020).
Studies show that monarch butterflies captively reared under insufficient conditions led
to weaker, smaller—which tend to impact flight ability—and more physically-pale
individuals, indicating inadequate captive rearing conditions (Tenger-Trolander et al.,
2019; Davis, Smith and Ballew, 2020). However, a study from 2021 shows that, when
35
using more accurate environmental targets and adding clarity to the captive rearing
procedures, migratory orientation was relatively unimpacted upon release of monarch
butterflies compared to other studies (Wilcox et al., 2021). This success shows promise
for captive rearing practices, though standardizing these practices becomes ambitious in
the case of Danaus plexippus, given the public participation in captive rearing.
Often long-standing captive rearing projects are considered successful in that they
prevent the demise of an at-risk species and, in some cases, re-establish populations,
however, species infrequently return from a critical point due to captive rearing that no
longer requires any intervention and supplementation. An ex-situ conservation effort
initiated in 1994 with the Karner blue butterfly (Lycaeides melissa samuelis) found
exceptional success in producing fit individuals compared to their wild
counterparts (Herms et al., 1996). Rearing of these butterflies occurred within
environmental chambers in a lab setting that achieved optimal environmental conditions
by imitating the species’ wild conditions; chambers maintained temperatures from 24º 26ºC with an 18:6 hour light-dark period and humidity at 57-68% (Herms et al., 1996;
Webb, 2010). After initial success, established propagation procedures include these
valuable environmental guidelines.
On a wildlife refuge in New Hampshire, reports showed less than 50 Karner blue
butterflies in the early 2000s, with one location becoming completely extirpated (Pau and
Holman, 2019). Through habitat restoration and population augmentation using proven
captive rearing methods, this site was reestablished in addition to other sites becoming
fortified, meeting the population goal of over 3000 individuals in a 2018 survey and
qualifying the species for delisting (Pau and Holman, 2019). While this species does still
36
require maintenance in the form of habitat restoration and monitoring and has not yet
been removed from the state or federal endangered species list, captive rearing did
eliminate the immediate risk of extinction and allowed for more time to research
ecosystem and life history requirements, likely contributing to the successful outcome of
this program and others for this species.
The listing status varied throughout the range of another important flagship and
indicator species, the Apollo butterfly (Parnassius apollo), with different European
countries experiencing different rates of loss initiating in the early 20th century
(Pierzynowska, Skowron Volponi and Węgrzyn, 2019). Regional extirpation of this
species resulted in total extinction in a number of countries, and the species, and the
Convention on International Trade in Endangered Species considers Apollo butterflies
near-threatened throughout their range in Europe (CITES, 2021). However, the IUCN
Red List previously listed the Apollo butterfly as near threatened throughout its entire
range, and as of 2021, the IUCN downgraded this status to least concern, though still
considered declining (Nadler et al., 2021). Delisting likely occurred due to the various
recovery actions taken throughout Europe, including ex-situ conservation methods.
One example of implementing these practices includes an exceedingly diminished
population in the Pieniny National Park in Poland; captive rearing significantly assisted
in increasing a critically at-risk population of 30-50 Apollo butterflies into a
metapopulation of 100-1200 individuals over 12 years (Adamski and Witkowski, 2007;
Pierzynowska, Skowron Volponi and Węgrzyn, 2019). However, initial trials of captive
rearing saw high mortality, and later witnessed adults eclosing with malformed or
sometimes missing wings (Witkowski and Adamski, 1996; Adamski and Witkowski,
37
1999; Pierzynowska, Skowron Volponi and Węgrzyn, 2019). Though this phenomenon
occurred in the wild, Witkowski and Adamski (1999) observed this at a higher rate in
female Apollo butterflies in captivity, instead of almost exclusively observed in males in
the wild.
Recent research seems to indicate that these malformations correlate with several
factors: genetic bottleneck effect leading to mutations, the insufficient chemical make-up
of the host plant, lack of exposure to an intracellular symbiont, or bacterial infection
(Łukasiewicz and Węgrzyn, 2015; Łukasiewicz, Sanak and Węgrzyn, 2016a, 2016b;
Pierzynowska, Skowron Volponi and Węgrzyn, 2019). Some describe that this
information could better inform captive rearing procedures to increase ex-situ success,
while others attempt to shift focus towards habitat corridors and wild translocation to
bolster the populations genetic integrity (Adamski and Witkowski, 2007; Dabrowski,
2008; Fred and Brommer, 2015). Though there appears to have been a trial-and-error
period to ascertain best practices for captive rearing of the Apollo butterfly, the ex-situ
program seemingly achieved great success in increasing population numbers even with
these trying aspects. This metapopulation allows for the opportunity to shift towards insitu conservation methods without immediate risk of extirpation.
Environmental Conditions & Captive Rearing
Staunchly contingent on temperature, development times and body size tend to
decrease in ectotherms as temperature increases until a certain point at which mortality
will likely occur if temperatures become extreme (Atkinson, 1994; van der Have and de
Jong, 1996; Angilletta, Steury and Sears, 2004; Fischer and Karl, 2010). Studies indicate
environmental conditions experienced in captivity contribute to morphological
38
discrepancies within captively reared butterflies, and as a notably influential factor of
development times, it could lead to severe survival repercussions (Nicholls and Pullin,
2000; Woodworth et al., 2002). Changes in environmental conditions encountered during
developmental stages may trigger phenotypic responses, expressed as variation in
proportions, especially if conditions continue to diverge through other life stages and
generations with the potential to act as a prominent selective pressure (Atkinson, 1994;
Loeschcke, Bundgaard and Barker, 2000; Steigenga and Fischer, 2009).
Contingent on the relationship between environmental conditions, development
times, productivity, and survival, the temperature-size rule supports findings of larger
body sizes at lower temperatures (Atkinson, 1994; Angilletta, Steury and Sears, 2004).
Morphological changes in butterflies explicitly impact an individual’s ability to fly,
consequently affecting the ability to properly disperse adequately and successfully
reproduce, in addition to diminishing competitive capability and fecundity (Berwaerts,
Van Dyck and Aerts, 2002; Norberg and Leimar, 2002). However, this rule cannot be
generally applied to ectotherms, and discerning whether this is a genetic or phenotypic
response to temperature depends on the species and requires research to determine
(Loeschcke, Bundgaard and Barker, 2000; Angilletta, Jr., and Dunham, 2003). Variation
in environmental conditions during captive rearing tend to have a reduced impact on
species living with greater environmental stochasticity; this can pose a challenge for more
specialized species, though insects often possess a certain margin of safety against
stochastic events (Inchausti and Halley, 2003; Crone, Pickering and Schultz, 2007).
A number of investigations into acclimation capacity uphold an optimal
temperature development hypothesis that indicates a threshold of moderate temperatures
39
produce more successful individuals, while success likely decreases outside of this
moderate temperature threshold (Huey and Berrigan, 1996; Huey et al., 1999; Woods and
Harrison, 2002). Moderate environmental conditions that mimic wild conditions without
extreme weather events that can occur out in the wild often become the goal when
captively rearing wildlife. Challenges can arise when these optimal environmental
conditions are not well understood for a species.
The endangered Puget blue butterfly, Icarcia icarioides blacmorei, also found in
WA South Puget Sound prairies, requires captive rearing to recover to prevent extinction.
Wild individuals were collected over two generations for rearing in captivity (Schultz,
Dzurisin and Russell, 2008). Studies compared survivorship, development rates, sex
ratio, biomass, size, and adult morphology between individuals reared in differing captive
circumstances to identify which conditions appropriately mimic wild conditions. To
determine this, the study then compared the morphology of captively reared adults to
wild adults to understand if divergencies occurred under any of the three captive rearing
conditions (Schultz, Dzurisin and Russell, 2008). One experimental group of Puget blue
butterflies was kept in refrigerators at the Oregon Zoo, a standard practice for other
butterfly rearing programs. The second group and third group, held at Washington State
University-Vancouver, were either reared in outdoor enclosures experiencing relatively
ambient conditions reared indoors in diapause chambers and experiencing temperature,
humidity, and light structures in a way to mimic best judgments of optimal conditions
(Dzurisin, 2005; Schultz, Dzurisin and Russell, 2008).
Results show survival and sex ratios were similar across all captive sites, and
other findings were consistent with the results of a long-term study that shows captive-
40
bred cabbage butterflies (Pieris brasicae) developed into smaller individuals than their
wild counterparts (Lewis and Thomas, 2001; Schultz, Dzurisin and Russell, 2008). These
findings are comparable to initial conversations with Mary Linders regarding the start of
the E. e. taylori captive rearing program where captively reared individuals appeared
smaller than wild individuals (Schultz, Dzurisin and Russell, 2008). However, reports
indicate initial success with the E. e. taylori captive rearing program at the Oregon Zoo
and continued success since the program expanded to Mission Creek Corrections Center
for Women, and these trials helped to determine life-history traits and develop
appropriate captive rearing methods and conditions utilized today (Grosboll, 2004;
Linders, 2007, 2011).
Depending on how long this trial-and-error period takes to determine appropriate
captive rearing conditions, risks imply that captive rearing as a primary method for
increasing population abundance may not always lead to enriching populations with fit
individuals (Lewis and Thomas, 2001; Norberg and Leimar, 2002; Schultz, Dzurisin and
Russell, 2008). If captive rearing must take place, one of the most critical factors to
ensure success is to mimic wild conditions in whatever way possible. Preferred practices
often include the use of controlled environmental chambers, which have seen continued
success at producing butterflies that are fit compared to their wild counterparts (Herms et
al., 1996). Studies found that outdoor enclosures during diapause allow environmental
conditions to mimic wild conditions and minimize the risks of selection pressures
changing because of divergent captive conditions (Nicholls and Pullin, 2000; Lewis and
Thomas, 2001).
41
To ensure selection is not occurring and promote the longevity of these successful
programs, comparisons and assessments between captive and wild individuals and their
environmental conditions must be carried out regularly to ensure they develop similarly.
Recommended methods of assessment include tacking morphological data which could
help reveal population disparities (Crone, Pickering and Schultz, 2007; Schultz, Dzurisin
and Russell, 2008). Assessments become especially important as the climate changes at
an increasing pace. However, research indicates high survival and productivity in
captivity leading to large-scale reintroductions may not augment populations until any
potential genetic and habitat concerns—including restoration and improving connectivity
to other populations— are addressed, and reintroductions occur during optimal weather
conditions (Oates and Warren, 1990; Lewis and Thomas, 2001; Nieminen et al., 2001;
Hanski, Ehrlich, et al., 2004).
42
METHODS
CAPTIVE REARING PROCESS
Butterfly Technician Environmental Management
Captive rearing at MCCCW is carried out in a 7.3 x 3.1 m glass greenhouse,
called greenhouse Raven (Figure 1). The design of this captive rearing greenhouse is
expected to provide relatively ambient environmental conditions while minimizing
extremes. There are two rooms in Raven—one smaller room (3 x 2.4 m) and one larger
room (3 x 4.9 m)—that can be partitioned by a glass door, the windows of which can
open and allow airflow between the two rooms. Raven has UV-transmitting glass panels
to allow for maximum sunlight and heat in addition to heaters attached overhead either
entrance to prevent freezing in the winter or help maintain environmental targets for
larvae in the spring. To avoid overheating in the summer, cooling is carried out by ceiling
exhaust fans, motorized dampers, and automated roof windows scheduled to open and
close, allowing passive ventilation when temperatures go above 86°F.
Two knit aluminum 50% reflective shade cloths cover the top of the greenhouse
during the summer months to prevent extreme heat caused by excessive sunlight
exposure. Butterfly Technicians Rearing Specialists monitor and maintain environmental
parameters within the greenhouse to reach environmental targets during each life stage by
increasing or decreasing the heat or fans, hanging sheets to block intense sunlight, using
humidifiers, and using icepacks, among other tactics. Technicians monitor environmental
conditions based on environmental targets (Table 1) using classic and digital
thermometers and relative humidity gauges showing minimum, maximum, and current
43
environmental conditions. Technicians record minimum and maximum temperature and
relative humidity daily onto a data form and graph this data compared to the targets.
A majority of the diapause stage occurs outside the greenhouse, in an 8’ x 10’ x 8’
shed with three 2’ x 2’ windows (Figure 1) and plenty of ventilation that allows for
ambient but not extreme environmental conditions. The shed has eight 4” x 10” grille
vents—six along the bottoms of three walls and two on opposing walls along the edge of
the ceiling—four 4” circular screen vents along the peak of the ceiling, and six panels
along the midpoint of the 12 m walls, with three 1” circular screen vents per panel.
Minimal maintenance is required of the Technicians to manage environmental conditions
during cold diapause.
Figure 1. MCCCW captive rearing greenhouse (left, photo credit: former Butterfly Program Coordinator) and cold
diapause shed (right).
Collecting Wild Adults
Collection of wild Euphydryas editha taylori adults occur from mid-April to early
May, although this period can vary by 2-4 weeks based on rain and other weather patterns
of that year (Linders et al., 2019). Usually occurs on Range 76 located at JBLM,
44
biologists search for and collect adults in the morning hours during the peak of E. e.
taylori flight season. Once captured, adults are placed in insect jars with proper
ventilation. The insect jars are then stored in small coolers (Figure 2) until they can be
transported to the captive rearing facility at MCCCW and processed by the Butterfly
Technicians.
Figure 2. Pictured are the containers in which wild females are transported to MCCCW, inside a cooler for
transport (left) and a wild female feeding (right).
Butterfly Technician Daily Care Procedures
All E. e. taylori daily care practices follow established husbandry manual
protocols developed by Oregon Zoo in 2009 for captive rearing and propagation (Barclay
et al., 2009). Specific captive rearing procedures for MCCCW have been created based
on approved protocols from the husbandry manual and refined over the years to
accommodate the differences in captive rearing facilities between the Oregon Zoo’s lab
and the MCCCW’s greenhouses (Curry et al., 2020). Procedures are currently in place
for E. e. taylori’s life cycle in captivity: oviposition, eggs to third instar, third instar to
fifth instar, warm diapause, cold diapause, wake up, post diapause, and release (Curry et
45
al., 2020) while older renditions of the procedures include pupation, adult eclosion,
captive females and males, and breeding. Note that captive breeding ceases in 2020 and
no longer take place at MCCCW. The goal the program is to produce at least 5,000 eggs
total that will result in 2,500 postdiapause larvae (Appendix A. Table 15).
Incarcerated Butterfly Technicians receive training from program partners prior to
the active rearing season to follow these protocols and procedures with minimal
supervision. Butterfly Program Coordinator visits the program a minimum of once a
week during the active rearing season to gauge program progress and facilitate
communication between partners. Technicians must be escorted out of the prison fence
around 0900 to carry out daily care tasks and remain there until they are brought back
inside around 1300, depending on the time of year.
Captive males were kept in large, mesh tents with several “brothers” and had
access to nectar plants, while captive females were stored in 16 oz deli cups inside a
refrigerator until breeding introductions occurred. Introductions were carried out in
portable popup tents, and once copulation occurred, Technicians moved females to an
oviposition chamber (Figure 3), and males retired to their large mesh tents. Lineages
were recorded using matrilines and patrilines to keep track of introductions and
copulations.
Technicians process the wild females by creating a matriline ID and placing them
into an oviposition chamber. Chambers are designed around a one-gallon potted Plantago
lanceolata, where they have access to a cotton ball soaked in a honey-water solution and
a water-soaked sponge. Technicians search oviposition chambers for eggs daily while
females have the opportunity to feed under a 16oz deli cup.
46
Figure 3. Females inside oviposition chamber, on the mesh closure (left), laying eggs (middle), and feeding on
honey-water solution (right).
If eggs are found, Technicians cut the leaf from the plant, remove as much of the
excess leaf as possible, perform and record egg estimates, and place eggs in a prepared
five oz deli cup lined with a folded paper towel. Females have the opportunity to oviposit
until they expire. Eggs and larvae are tracked and stored based on the matriline and cup
IDs. Eggs develop over 10-14 days, darkening from yellow to brown to purple, at which
point Technicians begin placing small leaves inside the cups preparing for eggs to hatch.
Eggs will hatch into first instar, and Technicians give larvae an increasing number of
leaves and replace paper towel liners as needed as larvae develop through first and
second instar (Figure 4).
Figure 4. First instar larvae at hatch (left), third instar larvae (middle), and egg and larval cup set-up (right).
47
Once larvae reach third instar, Technicians officially count and record the first
hard larval count of the season; Technicians count about 15 larvae per cup into 16 oz deli
cups lined with paper towels (Figure 4). Technicians continue prediapause larval care
procedures as larvae develop through third and fourth instar. Once larvae reach fifth
instar, warm diapause has officially started. About two weeks after allocation,
Technicians count and allocate larvae into 16 oz cups of 50 larvae per cup in-between
folded paper towels (Figure 5). Around mid-September, Technicians move larval cups
from the greenhouse outside to the diapause shed, and larval cups are stored on 12” terra
cotta dishes under 10” terra cotta pots with the drainage hole plugged with a cloth. Larvae
remain in cold diapause requiring minimal maintenance until mid-February.
Figure 5. Diapausing larvae (left) and cold diapause shed set up (right).
Wake-up occurs when the Technicians report high percentages of larval
movement following prolonged days of warmth. Larvae are brought into the greenhouses
and, Technicians allocate them into 16 oz release cups with about 15 larvae each. Most
48
postdiapause larvae will be released, which occurs two-three weeks after wake up,
depending on what the partner Biologist observes in the field. Retained larvae have the
opportunity to develop into sixth instar, where most pupate (Figure 6), but some will
reenter diapause, depending on environmental conditions. Pupae are stored in five oz deli
cups with a mesh covering until adults eclose about two-six weeks after pupation, at
which point Technicians record morphometric data and employ captive adult care
procedures.
Figure 6. 6th instar larvae (left) and pupae (right). Photo credit: Keegan Curry.
CAPTIVE REARING DATA COLLECTION
Productivity Outcomes Data Collection & Reporting
Captive rearing data—including breeding introduction data, oviposition data, egg
estimates, hatch rates, development rates and dates, number of larvae into and out of
diapause, and number of larvae to release—are recorded on a variety of data forms by
Butterfly Technicians (Appendix B, Figure 10). While continuing to capture the same
necessary information for reporting, these data collection forms were simplified by
49
Keegan Curry in 2019. This thesis used data from seven seasons, 2013-2014 through
2020-21. Note that the postdiapause larvae life stage was broken down into three
different groups for this study:
1. Postdiapause to Release
2. Postdiapause to Second Diapause
3. Postdiapause to Pupation
Environmental Data Collection
Honest Observer by Onset (HOBO) loggers are external data loggers that record
temperature, relative humidity, and light intensity at various intervals. These HOBO data
loggers are used to record relative humidity and temperature within the MCCCW E. e.
taylori captive propagation facility. These are placed in the same or similar settings as E.
e. taylori at various life stages to monitor environmental conditions during captive
rearing, see photos (Barclay et al., 2009). The life cycle for environmental conditions is
broken into six stages (Table 1).
HOBO loggers were programmed by connecting them to the Onset HOBOware
software using a USB cable and selecting launch logger. They are then programmed to
collect temperature and percent humidity every 1-3 hours, in addition to programing them
to collect minimum, maximum, and average temperature and humidity once daily. Hourly
data is used to calculate daily min/max/avg daily environmental data for seasons where
loggers were not programmed to collect daily data, or we are missing some of this daily
data. Historically, these HOBO loggers were deployed at the beginning of the season or a
new life stage and remained deployed until the start of another life stage or longer.
50
To keep environmental logging continuous, new loggers are brought in to
MCCCW to replace existing loggers, and previously deployed loggers are brought back
to The Evergreen State College and read using HOBOware software. The data is exported
to a CSV file then saved as an Excel workbook. Workbooks from the individual launches
are combined and organized into life stages by date for that season. The Butterfly
Program Coordinator summarizes each life stage’s minimum, maximum, and average
environmental data in Table 2. These summaries are compared to captive rearing targets
in Table 1 and reported on annually by SPP and WDFW to understand program success.
Table 1. Euphydryas editha taylori captive propagation environmental targets for all life stages.
Target
Life stage
Temp (°F)
%RH
50°-85°
Males
Females &
Oviposition
Eggs &
Prediapause
78°-85° for 2-6 h/day
≥50%
50°-90°
78°-90° for 4-8 h/day
≥50%
50°-90°
avg min ≤65°
≤65%
Warm Diapause
50°-90°
≥45%
Cold Diapause
≤35° for ≥60 days
≥50%
avg max
≤90%
≤45° night
Post- Diapause
≥65° daytime
>50° night
Pupation
≥65° daytime
≥50%
≥65%
51
Morphometric Data Collection
Butterfly Technicians measured adult butterflies’ weight (g) after captive eclosion
occurred or after wild females were delivered. Technicians placed butterflies inside an
enclosed scale and measured to the nearest 0.0001 gram (Figure 7). Weights were
recorded by Butterfly Technicians, then transcribed and summarized in Excel by the
Butterfly Program Coordinator for the annual report. Technicians measured butterfly
wing area (cm2) by delicately placing butterflies with their left side up inside a petri dish
and setting the dish on a 4 x 4 graphing paper. The butterflies ID tag is placed near it and
was photographed and used later to identify and double-check measurements. Butterfly
technicians recorded measurements, then the Program Coordinator transcribed and
summarized this data for annual reporting (Appendix A. Table 24). Wild populations had
smaller sample sizes since fewer wild females were available for measurements. Note
that morphometric data collection ceased in 2020.
Figure 7. Wing area measurements (left) and weight measurements (right) for adult females. Photo
credit: former Butterfly Program Coordinator.
52
DATA ANALYSIS
Productivity Data
Data is collected per matriline or larval ID, with a form created for every matriline
produced and brought into the greenhouse. Data forms inform each season’s annual
report to understand program success based on productivity targets; historically, captive
rearing targets at MCCCW have been for captive and wild females to jointly produce a
total of 2,500 postdiapause larvae from up to 5,000 eggs (Hamilton et al., 2013; Curry et
al., 2018). In the 2018-2019 season, this target changed to producing 1,800 postdiapause
larvae from up to 5,000 eggs (Curry et al., 2019). Reporting data was organized by life
stage to find where I used this data on E. e. taylori in captivity to find percent
productivity and survival from one life stage to the next, as follows:
1. Percentage of productive males were found by dividing successful copulations by
the total number of introductions that occurred that season.
2. Percentage of productive females were found by dividing the number of females
that have successfully oviposited by the total number of females brought to the
rearing facility that season.
3. Hatch rate was found by dividing the first hard larval count, executed once all
larvae have reached 3rd instar (prediapause count), divided by egg estimates at
oviposition.
4. Percent into diapause was found by dividing the number of larvae counted into
cold diapause cups by the first hard count of larvae at 3rd instar.
5. Percent out of diapause was found by dividing the larvae count out of diapause at
wake up by the number of larvae allocated to cold diapause.
53
6. Percent to release or retainment was found by dividing the total number of
surviving larvae at the end of the season by the number of larvae at wake up from
cold diapause.
7. Percent return to diapause was found by dividing the number of larvae that
reentered diapause by the number of retained larvae.
8. Percent to pupation was found by dividing the number of larvae that pupated by
the number of retained larvae.
9. Percentage of successful pupations and eclosions was found by subtracting the
number of unsuccessful eclosions from the number of pupations, then dividing
that number by the number of pupations.
Environmental Data
Hourly environmental data—organized by season and life stage—was accessed
from the SPP server, with 2013-2014 being the first season HOBO loggers were
consistently used to collect hourly temperature and humidity data. The 2019-20 season
was excluded from this study due to missing environmental data. Hourly data was
rounded to the nearest whole number and was then used to calculate daily minimum,
maximum, and average temperature and humidity for seasons or life stages where loggers
weren’t programmed to collect or were missing some of this daily data. Minimum,
maximum, and average environmental data was then summarized for reporting (Table 2).
These summaries for each season were averaged by life stage to produce an overall
average and overall range that was compared to the environmental targets, providing an
idea of environmental conditions experienced for all the seasons in this study (Appendix
A. Tables 16-22),
54
Rounded hourly data was compared to the targets using IF statements in Excel to
determine if they were within the target range or above/below the absolute target. When
looking to see if hourly data ever met the temperature targets during the eggs & prediapause life stage, targets for this life stage are temperatures between 50° to 90° for 90%
of the day. After determining whether a day was within the life stages environmental
parameters, a one or zero was used to indicate whether that day met the target (1) or not
(0). Ones were counted and divided by the number of days in that life stage for that
season, then converted to a percent. This was used as the percent of time targets were
met, with high percentages meaning targets were more frequently met.
Minimum and maximum daily temperatures were used to determine the percent of
days outside of temperature targets. IF statements were used to detect if temperatures in a
day went below or above absolute temperature targets by comparing the absolute
minimum temperature target to the minimum daily rearing temperatures and the absolute
maximum temperature target to the daily maximum rearing temperatures (Appendix B
Figure 11). This was done for all life stages except cold diapause, which is the only stage
without a temperature target range. The number of days outside the temperature targets
was summed and divided by the total number of days in the season to find the percent
outside the temperature target. For this methodology, high percentages mean targets were
less frequently met.
55
Table 2. Example summary of overall minimum, maximum, and average temperature (ºF) and relative
humidity (%) during captive rearing. This summary is produced for every life stage in each season.
Min
Max Avg
Min Temp
49º
59º
53º
Max Temp
69º
89º
81º
Avg Temp
57º
69º
64º
Min RH
26% 57% 38%
Max RH
66% 98% 82%
Avg RH
48% 81% 63%
Spearman’s Rho
Spearman’s rho (a non-parametric correlation coefficient) was calculated in JMP
Pro (version 16.1.0). Spearman’s rho was calculated between percent productivity and
survival rates and the percentage of time the relative humidity target was met, and
between percent productivity and survival and the percentage of time the temperature
target was met. This was done for all life stages in both wild and captive populations for
all seasons in this study. An alpha of 0.10 was used for statistical significance, to add to
the power of statistical tests, and p-values between 0.05 and 0.10 are pointed out
distinctly for the reader.
Morphological Data Analysis
Morphological data from the SPP server included 2013-14 to 2018-19 for weight,
and 2014-15 to 2018-19 for wing area. Only data on wild and captive female adults were
used. Raw data from area season was organized categorically depending on if the female
was wild or captive bred. This data was input in JMP and the Wilcoxon rank-sum test
56
was used to determine if the captive and wild morphometric data differed in each season.
Wilcoxon rank-sum was used to keep the analysis uniform because some seasons’
distributions were not normally distributed. Each season, wing area has smaller sample
sizes because individuals with damaged wings are excluded from the wing area
measurement.
In addition, in JMP a multiple linear regression was run using weight as a
response variable, and separately using wing area as a response variable, with both
captivity status (captive v. wild) and season (year) as predictor variables. In both cases
the overall model was statistically significant (for weight, F2,730 = 105.9, p < 0.001, for
wing area, F2,342 = 20.7, p < 0.001) albeit with relatively low adjusted R2 values (0.22 for
weight and 0.10 for wing area).
57
58
RESULTS
ENVIRONMENTAL DATA
Overview
Overall averages and average ranges summarize actual captive rearing conditions
at MCCCW for 2013-2020 (Table 3). Overall average temperature meets the respective
targets for males, females, prediapause, warm diapause, and pupation for nighttime.
Overall average relative humidity meets the targets for every life stage. Overall average
ranges are often outside of temperature target ranges and often meet relative humidity
targets.
Temperature
No life stages consistently met the targets 100% of the time (Table 4); pupation
met nighttime temperature targets 100% for 2/6 seasons and prediapause and warm
diapause for 1/7 seasons. Eggs and prediapause (4/7) and warm diapause (3/7) often met
the target 81-99% of the time. A majority of the targets are completed between 1-20% of
the time; males (2/6), females & oviposition (4/7), cold diapause (7/7), postdiapause (3/7
for nighttime, 4/7 for daytime), and pupation (2/6 daytime) having multiple seasons in
this range. Postdiapause (4/7 nighttime, 1/7 daytime) and pupation (2/6 daytime) both
have seasons where the target is never met (0%) (Appendix B. Figures 12-19).
59
Table 3. Overall averages and average ranges of environmental conditions compared to life stage targets for all life
stages in seasons 2013-2014 to 2018-2019 and 2020-2021.
Life Stage
Temp(°F)
Targets
(abs)
(50°-85°)
Males
78°-85° for 26 h/day
Females &
Oviposition
Eggs &
Prediapause
(50°-90°)
78°-90° for
4-8 h/day
(50°-90°)
avg min ≤65°
Overall Average
(Overall Avg
Range)
64
(47-92)
Cold Diapause
≤45° night
Postdiapause
≥65° daytime
>50° night
Pupation
≥65° daytime
≥50%
65
Overall Average
(Overall Avg
Range)
60
(27-89)
62
≥50%
(46-94)
(27-88)
67
64
≤65%
(48-97)
(29-91)
70
65
≥45%
Warm Diapause (50°-90°)
≤35° for ≥60
days
%RH
Targets
(52-97)
(28-91)
44
86
(19-84)
58
≥50%
≥65%
(40-94)
68
(40-85)
(29-94)
62
69
≥50%
(47-90)
(31-95)
Relative Humidity
No life stages consistently met the targets 100% of the time (Table 4); warm
diapause and cold diapause met the relative humidity target 100% for 1/7 seasons each. A
majority of the targets are either met 21-40% of the time—males and postdiapause
(2/7)4/6), females & oviposition (3/7), eggs & prediapause (2/7), and pupation (2/6
60
daytime)—or met between 81-99% of the time; females (2/7), cold diapause (6/7), and
postdiapause (2/7). Eggs & prediapause (4/7) typically met the target 1-20% of the time.
No life stages never met the relative humidity target (0%) (Appendix B. Figures 12-19).
Table 4. Average and range of the percent of time the environmental targets are met for all life stages for
seasons 2013-2014 to 2018-2019 and 2020-2021.
Target
Life Stage
Males
Females &
Oviposition
Eggs &
Prediapause
Temp (°F)
%RH
(50°-85°)
78°-85° for
2-6 h/day
≥50%
50-90
(2%-67%)
(21%-93%)
18%
51%
(0%-48%)
(14%-95%)
85%
26%
(47%-100%)
85%
(1%-96%)
74%
(59%-100%)
(57%-100%)
7%
97%
(2%-16%)
(94%-100%)
≥50%
(50°-90°)
78°-90° for
4-8 h/day
≤65
avg min ≤65
Warm
Diapause
(50°-90°)
≤65%
avg min ≤65°
Cold
Diapause
≤35 for ≥60
days
PostDiapause
%Time Targets Met
Temp (°F)
%RH
Average
Average
(Range)
(Range)
34%
44%
≥45%
≤45° night
≥50%
>50 night
≤45° night
≥50%
Pupation
≥65° daytime
4%
(0%-15%)
21%
(0%-72%)
86%
(41%-100%)
16%
(0%-36%)
69%
(47%-88%)
36%
(8%-71%)
61
Outside Temperature Targets
Overall, the percent of time the minimum daily temperature is below the absolute
minimum temperature target for each life stage is relatively consistent from season to
season except for 2013-2014 (Table 5). Males (2/6), females (3/7), prediapause (3/7),
warm diapause (6/7), and pupation (2/6) were often never outside of the target. Males
(3/6), females (2/7), warm diapause (3/7), and pupation (2/6) were often only outside of
the target 1-21% of the time. Postdiapause (3/7) was often outside the target 81-99% of
the time. During the 2018-2019 season, postdiapause was always outside the nighttime
target (100%) (Appendix B. Figures 20-27) The minimum target was frequently greater
than the absolute minimum target, either 61-80% (3/7) or 81-99% (4/7) of the time.
Overall, the percent of time the maximum daily temperature is above the absolute
maximum temperature target for each life stage is relatively consistent from season to
season (Table 5). Males (1/6), females (2/7), warm diapause, postdiapause, and pupation
(1/7) were occasionally never outside of the target. However, males (2/6), females (4/6),
prediapause (6/7), warm diapause (3/7), postdiapause (4/7), and pupation (4/7) were often
only outside of the target 1-20% of the time. During the 2020-2021 season, pupation was
the only life stage to be between 81-99%. (Appendix B. Figures 19-26)
62
Table 5. Average and range of the percent of days outside of the environmental targets for all life stages for
seasons 2013-2014 to 2018-2019 and 2020-2021.
Target
Life stage
Males
Females &
Oviposition
Eggs &
Prediapause
Temp (°F)
% Outside Target
Below Target Above Target
Average
Average
(Range)
(Range)
(50°-85°)
19%
22%
78°-85° for 2-6
h/day
(0%-84%)
(0%-38%)
50-90
16%
12%
(50°-90°)
(0%-58%)
(0%-26%)
78°-90° for 4-8
h/day
7%
10%
(0%-40%)
1%
(2%-26%)
21%
(0%-8%)
(0%-53%)
avg min ≤65
Warm
Diapause
(50°-90°)
avg min ≤65°
Cold Diapause
≤35 for ≥60 days
(50°-90°)
--
≤45° night
53%
21%
≥65 daytime
(0%-100%)
(0%-69%)
≤35° for ≥60 days
18%
14%
≥65 daytime
(0%-59%)
(0%-50%)
Post- Diapause
Pupation
88%
(75%-97%)
63
PRODUCTIVITY & SURVIVAL DATA
Captive Population
Male productivity was consistently low until the 2018-19 season, when it peaked
at 62.7% (Table 6). Female productivity was at or above 90% for the first three seasons,
then dropped significantly for 25% in the 4th season, and remained below 75%
productivity for the last two seasons. Hatch rates vary greatly by season. Percent into
diapause remained above 80% for all six seasons, and percent out of diapause and percent
to release/retainment never went below 96%.
Table 6. Percentage of productivity & survival of captive E. e. taylori populations at MCCCW for seasons 2013-2014
to 2018-2019.
Season
% Males
% Females Hatch Rate % Into
Productive Productive
Diapause
% Out of
Diapause
% To
Release
2013-14
2014-15
2015-16
2016-17
2017-18
2018-19
Average
Range
10.6%
27.8%
16.0%
10.7%
7.7%
62.7%
23%
8%-63%
99.9%
99.6%
98.0%
100.0%
100.0%
96.7%
99.0%
97%-100%
99.8%
100.0%
99.8%
99.4%
100.0%
96.2%
99.2%
96%-100%
64
93.8%
95.2%
90.0%
25.0%
71.4%
66.7%
73.7%
25%-95%
62.0%
29.8%
33.3%
49.2%
24.8%
96.1%
49.2%
25%-96%
99.0%
94.6%
86.7%
98.2%
96.7%
97.1%
95.4%
87%-99%
Wild Population
Females’ productivity varied between 70-100% for all 7 seasons, with an average
of 85.5% (Tables 7 & 8). The hatch rate remained above 80%, going above 100% for 3
seasons since egg counts are estimates. Percent into diapause is its lowest in the 2014-15
seasons but stays above 97% for every other season. Percent out of diapause remained
above 96% for all seasons and percent to releases/retained remained above 98% for all
seasons. Return to diapause is highest from in the last three seasons (2013-2018), and
pupation is lowest in 2013-14, 2016-17, and 2017-18. Eclosion is consistently high, with
2015-2016 having the lowest percent success (79%).
65
Table 7. Productivity and survival of the wild E. e. taylori populations at MCCCW for adults to postdiapause
stages during seasons 2013-2014 to 2018-2019 and 2020-2021.
Season
% Females
Productive
Hatch Rate
% Into
Diapause
% Out of
Diapause
2013-14
2014-15
2015-16
2016-17
2017-18
2018-19
2020-2021
Average
Range
91.7%
75.0%
85.0%
70.0%
91.3%
100.0%
85.7%
86%
(70%-100%)
97.1%
92.5%
80.3%
114.4%
104.9%
101.2%
95.2%
98%
(80%-114%)
99.5%
90.4%
98.2%
99.2%
98.1%
97.1%
99.0%
97%
(90%-100%)
99.8%
98.1%
96.6%
99.7%
99.8%
99.8%
99.9%
99%
(97%-100%)
Table 8. Productivity and survival of the wild E. e taylori populations at MCCCW for release/retainment to
eclosion stages during seasons 2013-2014 to 2018-2019 and 2020-2021.
66
% Successful
Pupation &
Eclosion
58.0%
92.0%
76.3%
88.5%
85.4%
78.5%
52.9%
86.8%
57.6%
93.6%
60.6%
87.2%
--65%
88%
(53%-85%) (79%-94%)
Season
% Released/
Retained
% Return to % Wild to
Diapause
Pupation
2013-14
2014-15
2015-16
2016-17
2017-18
2018-19
2020-2021
Average
Range
100.0%
100.0%
99.9%
98.9%
99.9%
99.1%
100.0%
100%
(99%-100%)
12.0%
8.3%
13.2%
37.4%
27.0%
25.1%
-20%
(8%-37%)
SPEARMAN’S RHO CORRELATION
Captive Population – Productivity/Survival vs. Environmental Targets
The percent success of different life stages for the captive population was not
typically correlated with the percentage of time the environmental targets were met.
Using Spearman’s rho, the percent of ovipositing females was positively correlated with
percentage of time %RH is meeting the target (rho = 0.77, p = 0.07, Table 9). However,
the percentage of successful ovipositing females and % of time temperature met the
target was negatively correlated (rho = -0.75, p = 0.08, Table 9). Also note that these pvalues were between 0.05 and 0.10 (Appendix B. Figures 28-33).
Table 9. Spearman’s rho coefficient for the percent productivity/survival of the captive population versus
percent of time the environmental targets were met for seasons 2013-2014 to 2018-2019 and 2020-2021.
Captive Population
Life Stage
Males &
Breeding
% Productivity
/Survival Rates
% Successful
Copulations
Spearman’s Rho Coefficient
% Time RH
Target is Met
% Time Temp
Target is Met
0.17
-0.08
Females &
Oviposition
Eggs &
Prediapause
% Females
Productive
0.77*
-0.75#
Hatch Rate
-0.26
-0.26
Warm Diapause
% Into Diapause
0.31
-0.20
Cold Diapause
% Out of Diapause
-0.25
0.07
Postdiapause
% From Wake Up
to Release
0.00
Daytime
Nighttime
-0.24
0.31
* p = 0.07
# p = 0.08
67
The percent success of different life stages for the captive population was not
typically correlated with the percentage of days outside the temperature targets. Using
Spearman’s rho, the percent of prediapause larvae was positively correlated with the
precent of days with temperatures below the minimum target (rho = -0.88, p = 0.02,
Table 10). However, the percent of successful ovipositing females was negatively
correlated to the percent of days with temperatures above the maximum target (rho = 0.78, p = 0.07, Table 10) in addition to the percent of larvae out of diapause being
negatively correlated to the percent of days with temperatures above the maximum target
(rho = -0.75, p = 0.08, Table 10). Also note that these p=values were between 0.05 and
0.10 (Appendix B. Figures 42-47).
Table 10. Spearman’s rho coefficient for the percent productivity/survival of the captive population versus
the percent of time outside the temperature targets for seasons 2013-2014 to 2018-2019 and 2020-2021.
Captive Population
Life Stage
Males
Females &
Oviposition
Eggs &
Prediapause
% Productivity
/Survival Rates
% Males
Productive
% Females
Productive
Hatch Rate
Warm Diapause % Into Diapause
Cold Diapause
% Out of Diapause
Postdiapause
% To Release
^ p = 0.07
*p = 0.02
68
Spearman’s Rho Coefficient
% Below Min
Temp Target
% Above Max
Temp Target
-0.66
-0.06
-0.17
-0.78*
0.88^
0.58
0.65
0.20
--
-0.46
-0.29
-0.09
Wild Population – Productivity/Survival vs. Environmental Targets
The percent success of different life stages for the wild population was not
typically correlated with the percentage of time the environmental targets were met.
Using Spearman’s rho, the percent of larvae that pupated was negatively correlated to the
percentage of time night temperature targets are met (rho = -0.76, p = 0.08, Table 11).
However, the percent of successful ovipositing females was positively correlated to the
percent of time outside the percent females productive (rho = 0.88, p = 0.02, Table 11)
Also note that these p=values were between 0.05 and 0.10. (Appendix B. Figures 34-41)
Table 11. Spearman’s rho correlation for the percent productivity/survival of the wild population versus the
percent of time the environmental targets were met for seasons 2013-2014 to 2018-2019 and 2020-2021.
Wild Population
Life Stage
Females &
Oviposition
Eggs &
Prediapause
Warm Diapause
Cold Diapause
Postdiapause to
Survival
Postdiapause to
2nd Diapause
Postdiapause to
Pupation
Pupation
Spearman’s Rho Coefficient
% Productivity
/Survival Rates
% Females
Productive
Hatch Rate
% Into
Diapause
% Out of
Diapause
% To Release
or Retained
% Wilds Return
to 2nd Diapause
% Wilds to
Pupation
% Wilds
Enclosed
% Time RH
Target is Met
% Time Temp Target
is Met
-0.25
0.23
-0.11
-0.44
0.39
0.07
-0.68^
-0.28
0.22
-0.03
-0.29
0.31
Daytime
-0.12
Daytime
-0.20
Daytime
-0.12
Daytime
-0.12
Nighttime
-0.11
Nighttime
0.39
Nighttime
-0.76*
Nighttime
-0.09
^ p=0.09
* p=0.08
69
No correlations occurred for any life stages in the wild population compared to
the percent of days outside the temperature targets (Table 12; Appendix B. Figures 4855).
Table 12. Spearman’s rho coefficient between the productivity/survival of the wild population versus the
percent of time outside the temperature target for seasons 2013-2014 to 2018-2019 and 2020-2021.
Wild Population
70
Spearman’s Rho Coefficient
Life Stage
% Productivity
/Survival Rates
% Below Min
Temp Target
% Above Max
Temp Target
Females &
Oviposition
% Females
Productive
0.04
0.13
Eggs &
Prediapause
Hatch Rate
0.19
-0.19
Warm Diapause
% Into
Diapause
0.61
-0.04
Cold Diapause
% Out of
Diapause
--
-0.44
Postdiapause to
Survival
% To Release
or Retained
-0.26
0.26
Postdiapause to 2nd
Diapause
% Wilds
Return to 2nd
Diapause
0.03
0.09
Postdiapause to
Pupation
% Wilds to
Pupation
0.26
-0.31
Pupation
% Wilds
Successfully
Enclosed
-0.06
-0.26
MOPHOLOGICAL RESULTS
Weight & Wing Area
Captively bred adult females weighed significantly more than wild females in all
six seasons (Table 13, Figure 8, Appendix B Figures 56-61). There was not an effect of
season on butterfly weight (over time, Table 13). Differences in wing area between the
captive and wild females were more variable, as wild adults had significantly larger
wings during the 2014-2015 and 2017-2018 seasons but not others (Table 14, Figure 9,
Appendix B. Figures 62-66). Both captivity status (wild vs. captive) and season had
significant effects on wing area (Table 14), with a slight decrease in wing area across the
seasons (visible in Figure 9).
Table 13. Median adult female E. e. taylori weights by season at MCCCW, with p-values from a Wilcoxon
rank sum test (see also Appendix B Figures 36-61).
Median Weight (g)
Season
Captive
Wild
2013-14
2014-15
2015-16
2016-17
2017-18
2018-19
0.18
0.15
0.17
0.17
0.15
0.17
0.14
0.09
0.11
0.12
0.12
0.12
Multiple Regression on Weight
p-value
Parameter
Estimate
p-value
0.03
<0.0001 Status[Captive]
0.022 <0.001
<0.0001
Season
-0.001 0.114
<0.0001
<0.0001
R2adj = 0.22
<0.0001
Table 14. Median adult female E. e. taylori wing area by season at MCCCW, with p-values from a Wilcoxon
rank sum test (see also Appendix B Figures 62-66 ).
Median Wing Area (cm2)
Season
Captive Wild p-value
2014-15
1.43 1.68 0.0118
2015-16
1.62 1.61 0.5535
2016-17
1.51 1.63 0.4588
2017-18
1.34 1.51 0.0035
2018-19
1.24 1.28
0.273
Multiple Regression on Area
Parameter
Estimate
Status[Captive]
Season
-0.037
-0.065
p-value
0.011
<0.001
R2adj = 0.10
71
Figure 8. Data points and boxplots of adult female weight (g), by captive (black points) or wild (grey points) status and
across the 2013-14 to 2018-19 seasons (x-axis labels). The light pink boxes represent 10th and 90th percentiles, the red line
the mean, and the reddish boxes the 95% CI around the mean.
Figure 9. Data points and boxplots of adult female wing area (cm2), by captive (black points) or wild (grey points) status
and across the 2013-14 to 2018-19 seasons (x-axis labels). The light pink boxes represent 10th and 90th percentiles, the red
line the mean, and the reddish boxes the 95% CI around the mean.
72
DISCUSSION
Adult Butterflies
Both the percentage of time the environmental targets were met during different
stages of captive reading, and butterfly productivity and survival rates, were variable. For
different life stages, there was a lack of correlation between the percent productivity and
survival and the percent of time environmental targets were met (Table 9) in addition to a
lack of correlation to the percent of days outside the temperature targets (Table 10)
throughout the seasons in this study. Whether or not males successfully copulate (%
males productive) experienced variation throughout the seasons but was relatively low,
with average productivity of 23% (Table 6). Environmental conditions could influence
these outcomes, given the need for adult butterflies to bask for thermoregulation and
capacity to become exhausted by high heat (Porter, 1982; Weiss, Murphy and White,
1988; Hellmann et al., 2004). However, these factors lack any correlations in this study.
Females and oviposition success (% females productive) also experiences
variation, particularly in captive females yielding a range of 25%-95% productivity
(Table 6). The percent of captive females productive and the percent of time the
temperature target is between 78º-90º for two to six hours per day (Table 9) was
negatively correlated, in addition to a negative correlation to the percent of days the
maximum daily temperature was above the maximum target of 90º (Table 10). However,
no such correlations exist for wild females. Captive females could be experiencing
diminished environmental plasticity compared to their wild counterparts. However,
parameters often used to compare phenotypic plasticity between the wild and captive
73
populations would be challenging to collect for this program; ages and first-reproductions
of wild individuals are unknown, limiting insight into fecundity (Stillwell and Fox, 2005;
Steigenga and Fischer, 2009). Current research on fitness of captively reared and
captively bred E. e taylori has focused on high larval mortality experienced during
diapause and postdiapause at Oregon Zoo and investigating oviposition outcomes and
larval host plant preferences (Lewis et al., 2018; Haan, Bowers and Bakker, 2021).
Parameters available to investigate captive fitness within this program include
reintroducing morphometric data collection, adding egg measurements via microscopes,
recording and quantifying development times seasonally (Fischer, Brakefield and Zwaan,
2003; Stillwell and Fox, 2005; Crone, Pickering and Schultz, 2007).
Captive female wing areas were similar to or smaller than wild female wing area
(depending on the season), with wild wing area on a downward trend overall (Table 14,
Figure 9). For one of these seasons (2014-2015), the wild sample size was only seven
compared to the captive sample size of 29. Similarly, the 2016-2017 season, which did
not show a significant difference (Table 14), also had sample size disparities with the
wild sample size at ten versus the captive sample size of 124. Differences in sample sizes
between captive and wild populations always exist given the quantity of postdiapause
larvae retained and because morphometric data is recorded for all captive adults that
eclose even if they get released. This disparity could skew results, so a larger data pool,
including Oregon Zoo morphometric results, could help inform this research. These
results are consistent with research trialing optimal rearing conditions for the Puget blue
butterfly, which resulted in smaller wings and body lengths, and other earlier findings
74
showing similar outcomes (Lewis and Thomas, 2001; Schultz, Dzurisin and Russell,
2008; Altizer and Davis, 2009).
Captive and wild females’ weights were significantly different for all seasons
(Table 13), with captive females consistently being heavier than wild females (Figure 8).
This appears to be consistent with research that found larger or heavier body sizes in
captive butterflies (Lewis and Thomas, 2001; Schultz, Russell and Wynn, 2008) but
inconsistent with a study showing smaller body sizes in captive individuals (Schultz,
Dzurisin and Russell, 2008). Larger sizes do tend to be linked to lower temperatures
(Atkinson, 1994; Angilletta, Steury and Sears, 2004), and some studies indicate selective
pressure towards lower temperatures could decrease productivity during warmer
conditions (Berger, Walters and Gotthard, 2008; Fischer and Karl, 2010) however this is
not a general rule for all ectotherms (Loeschcke, Bundgaard and Barker, 2000; Angilletta,
Steury and Sears, 2004; Stillwell and Fox, 2005). Research also suggests a positive
relationship between temperature, body size, and fecundity (Blanckenhorn, 2000;
Gotthard, Berger and Walters, 2007; Berger, Walters and Gotthard, 2008) however the
outcomes of this limited study are not consistent with these findings. Studies of
temperature variation impacts on checkerspots and E. e taylori focus predominantly on
the relationship between larval development times and how that relates to host plant
availability and adult emergence timing (Weiss, Murphy and White, 1988; Weiss et al.,
1993; Hellmann et al., 2004; Bennett, Betts and Smith, 2014). In addition, some studies
comparing body size look at body area or length rather than body mass, making it unclear
if these are comparable.
75
Egg & Larval Stages
Captive hatch rates had greater variation than wild hatch rates (Tables 6 & 7).
However, there was a lack of correlation between prediapause survival and whether or
not the environmental targets were met (Tables 10 & 12). It’s unclear what caused the
captive population to have more variation in hatch rates than the wild population, given
that primary causes of egg loss in the wild are predation—which was not a factor in
captive rearing—or desiccation by extreme temperatures (Kuussaari et al., 2004).
However, egg clusters can be exceedingly challenging to count depending on how many
eggs were laid in a single cluster, so egg counts are considered estimates. Wild and
captive hatch rates would still see similar variation if this were the sole factor.
Prediapause (% into diapause), warm and cold diapause (% out of diapause), and
postdiapause (% to release/retained) life stages do not have a lot of variation between
seasons for either captive or wild populations (Tables 6-8). The success of these life
stages and the percentage of time environmental targets were met lack any correlation for
the captive population (Tables 10 & 12). However, a positive correlation was indicated
for the captive-bred population between eggs and prediapause larvae (hatch rate) and the
percent of days below the minimum temperature target of met (Table 11). Exceedingly
high temperatures can lead to egg desiccation and larval death, while low temperatures
slow development (Hellmann et al., 2004; Kuussaari et al., 2004). There has a positive
correlation between temperatures below 50° and prediapause larval survival for the
captive population (Table 11), which could support acclimatization to cool
conditions (Blanckenhorn, 2000). Seasons with the highest captive egg production, 20132014, 2016-2016, and 2018-2019, interestingly have the lowest minimum temperature,
76
with two of those seasons also having the highest maximum temperatures for all of the
seasons in this study (Appendix A. Tables 17-22), making these findings vague.
It’s unclear how the percentage of time environmental targets were or were not
met during prediapause larval stages impacts the timing of other life stages and how that
impacts success once released. Prediapause, postdiapause, and pupation development
times strongly rely on microclimate conditions. During any of these stages, development
times influence the timing of other life stages, which can impact adult emergence time, a
primary determinant of reproductive success (Weiss et al., 1987, 1993; Hellmann et al.,
2004; Kuussaari et al., 2004). Tests indicated a negative correlation exists for
postdiapause larvae to second diapause (% Return to diapause) and the percent of time
nighttime temperature target is met (Table 11). Postdiapause development is more
strongly reliant on the ability to thermoregulate within the microclimate the larvae are in
(Weiss, Murphy and White, 1988; Hellmann et al., 2004).Wild larvae also have a
negative correlation to the percent of time the humidity target being met during diapause;
however, the direct impacts of humidity on captive rearing are less understood given
humidity’s usual relation to host plant senescence in conjunction with temperature
(Hellmann et al., 2004; Klockmann and Fischer, 2019; Reed et al., 2019).
Program Success
Euphydryas editha taylori captive rearing seasons at MCCCW tend to be
successful in producing enough eggs that hatch larvae that survive through all life stages
in captivity and make it to be released in the wild (Tables 6-8). A 2016 periodic update
of E. e. taylori and their habitat showed a prosperous population on JBLM range 76, near
where MCCCW captively reared larvae are released (Potter, 2016). However, this
77
population was not observed during brief inspections in the 2021 E. e. taylori flight
season. Access to this site was restricted following the impacts of COVID-19 and official
surveys and reports have yet to officially confirm this population’s continued presence in
the site (pers. communication, Mary Linders). This development isn’t necessarily an
indication that captive rearing is producing unsuccessful individuals since bounteous
flight seasons have been observed on range 76 weeks after seemingly successful
translocations of captive populations from MCCCW to release sites have taken place.
Poor host plant availability and unprecedented weather events in the field are thought to
play a role in why butterflies were not observed this past year. Euphydryas editha
subspecies are also notorious for disappearing and reappearing depending on seasonal
success (Hanski, Ehrlich, et al., 2004; Hellmann et al., 2004). When compared to the
captive rearing program for the critically endangered Euphydryas editha quino—which
was able to release a total of 1,513 larvae over two captive rearing seasons (CBI Blog,
2020)—E. e. taylori program goals of harvesting 2,500 postdiapause larvae from 5000
eggs seem substantial (Appendix A. Table 16). MCCCW, though it has only passed the
threshold of 5000 eggs harvested once in the 2018-2019 season, exceeds the postdiapause
larvae target every season, excluding 2017-2018 and 2020-2021 (Appendix A Table 23).
When compared to the trials of inexplicable larval mortality experienced during diapause
and postdiapause at the Oregon Zoo, MCCCW captive rearing program was highly
prosperous; per the Zoos 2017-2018 report, 56% of the near 1500 larvae into diapause
survived to postdiapause, and 28% of the larvae that entered diapause survived to release
(Lewis et al., 2018).
78
The other remaining E. e. taylori captive rearing program in OR—separate from
this program and Oregon Zoo, and located in Coffee Creek Corrections Facility
(CCCF)—set a goal for 2020-2021 to exceed the previous season postdiapause release of
1100, exceeding that goal by 136 (Naseth, 2021). However, mortality experienced after
postdiapause was relatively high compared to MCCCW, with 66% survival from
diapause to release. For comparison, MCCCW experienced 90% survival from the start
of diapause to release for 2017-2018 and 99% survival for 2020-2021. The Zoo and
CCCF rear in a lab setting, excluding cold diapause where larvae are moved outside to an
overwintering area per research and protocol (Schultz, Dzurisin and Russell, 2008;
Barclay et al., 2009) However, providing a significant number of larvae for release alone
is not related to reintroduction success (Oates and Warren, 1990; Hanski, Ehrlich, et al.,
2004, p. 281). In the labs, wild conditions need to be replicated, as opposed to the
greenhouses at MCCCW, where near-ambient conditions need to be manipulated.
Comparisons could be carried out between these three facilities to fully understand
survival differences and the impacts of environmental conditions, respectively. Ongoing
efforts to improve habitat connectivity, remove invasive species, and restore prairies
strengthen chances of captive rearing success (Potter, 2016). Coupling habitat progress
and conditions with program outcomes and conditions is likely the next step in future
research that would provide the greatest benefit to future program, and species recovery,
success.
Recommendations to Environmental Targets & Procedures
It is imperative for those working with the Butterflies Technicians to focus on
instilling the importance of managing and recording the environmental conditions. Tying
79
the control of the environmental conditions firmly to the productivity and survival of the
butterfly brings this into the purview of E. e. taylori daily care needs, rather than an
isolated task that may appear nonessential or may become easily forgotten. Once the
Technicians receive the necessary training and educational background, they should
receive clear procedures dedicated to detailing the environmental targets for each life
stage and what the Technicians can do to adjust conditions and bring them closer to the
objective conditions. Studies show that when captive rearing procedures—including
environmental parameters—are detailed and comprehensive, there is a higher chance for
success (Adamski and Witkowski, 2007; Crone, Pickering and Schultz, 2007; Wilcox et
al., 2021). This empowerment through education allows Butterfly Technicians to work as
a team to command the decision-making for the greenhouse based on the tools available
to them. Rapid response by technicians could minimize loss of butterfly production or
larval mortality that may otherwise occur while waiting for communication between
Technicians and the Butterfly Program Coordinator.
Established parameters for maintaining environmental conditions were initiated
but could be expanded upon and developed into Captive Rearing Environmental Target
Procedures. An example could be females and oviposition, where the temperature should
be between 78º-90º for 4-8 hours of the day. This translates to moving the ovipositing
chambers into sunlight within the greenhouse but still providing the option of shade—
either through the plantain leaves or adding a sticky note to the top of the chamber—to
promote egg-laying but prevent overheating (pers. communication, Mary Linders). This
allows thermoregulation but prevents exhaustion and early mortality. In addition, there is
a very brief description in the husbandry manual explaining basking and the need to
80
provide a variety of light, temperature, and humidity microclimates for basking adults
and postdiapause larvae. Basking information has not been elaborated on in the MCCCW
procedures, making it unclear what basking behaviors to expect.
Photos throughout the procedures also show ceilings or shelves draped in sheets
during oviposition, eggs and early instar larvae, and postdiapause larvae life stages
(Barclay et al., 2009; Curry et al., 2020). This could be a tactic to prevent extreme heat,
or the set-up technicians use to block the intense setting sun before leaving for the day.
However, this is unclear and conflicts with the directive in the husbandry manual and the
basking behavior that this species is known to carry out during multiple life stages
(Weiss, Murphy and White, 1988). This information has been briefly described in captive
rearing procedures, but incorporating and situating this kind of information into
environmental procedures emphasizes the importance of environmental conditions during
captive rearing. Covering portions of the greenhouse with shade cloths allows for more
control and providing temperature thresholds in which these should be erected could be
imperative to productivity and development. Providing explicit instruction on how to
achieve—or get close to achieving—environmental targets in relation to the outcomes—
or consequences—of carrying out these tasks can prevent conditions from swinging from
one extreme to another.
Environmental targets for reporting and rearing could be slightly different to
prevent overreporting but still provide the Butterfly Technicians with explicit instructions
for managing environmental conditions. Life stages where targets are rarely met could
cause misalignment in the Butterfly Technicians’ understanding of why this is important
and lead to despondency at their work. Cold Diapause, as an example, has an average
81
mean temperature of 44 and an average minimum temperature of 19 (Table 3), with the
2016-2017 season having the highest percentage of time meeting the target at 16% (Table
4). This temperature target could be slightly increased since the lack of meeting the target
doesn’t appear to impact larval survival due to the lack of mortality out of diapause. This
is especially true since average temperatures in Belfair remain above 35 degrees for 11
months of a year (Appendix B. Figure 67) based on records available online (Weather
Spark, no date). An absolute minimum temperature could be applied to females and
oviposition—and males if breeding returned—so Technicians know to readily implement
adjustments to conditions if the overnight temperature dipped below the specified
absolute minimum. Recording and properly managing ambient conditions using HOBO
loggers at MCCCW could better inform this process.
Providing an absolute range for humidity during all life stages may allow
Technicians to better understand what steps they should take to adjust conditions and
prevent loss of production or larval mortality that may occur from females becoming
waterlogged or egg and larval cups becoming moldy due to high humidity. Currently,
there’s no indication that high humidity could be unfavorable with these targets. This lack
of clarity could prolong daily care when even clean larval cups need fresh liners due to
saturation by condensation. Providing an absolute range for humidity—such as removing
shoe bin lids, moving shelves near open windows and vents during the day, and ensuring
plantain leaves are thoroughly dry for all larval stages—could keep humidity closer to the
targets and reduce the amount of time Technicians spend on E. e. taylori daily care
during transition periods. Providing an absolute range for temperature during all life
82
stages allows for a similar response by technicians; the range indicates when action
should be taken to prevent temperatures from going to an extreme.
Once there is a better capacity for maintaining these targets, a qualitative aspect
could be added to the adult life stages in addition to the numeric temperature targets. The
Technicians are required to put adults in light for thermoregulation for a portion of the
day per protocols and procedures, seemingly indicated by the specificity of the targets
during those life stages in comparison to other targets (Boggs and Nieminen, 2004;
Barclay et al., 2009; Curry et al., 2020). Since breeding no longer occurs—when carrying
out and checking ups on copulation tents took up a significant amount of the Technicians’
time—maintaining these targets for ovipositing females could involve Technicians
regularly checking on activity in-between searching chambers for eggs. Technicians
could record behavioral observations, and productivity and environmental conditions will
be documented. Activity for the day could be summarized by the team and discussed with
the Butterfly Program Coordinator at the end of the life stage. This joining of qualitative
and quantitative data could inform annual reporting to understand how the environmental
conditions impacted productivity. The same could be done for males if breeding were to
return; qualitative notetaking was extensively carried out before breeding ceased, and
incorporating this portion back in could greatly assist in improving program
communication. This quantitative and qualitative data could be used to determine if a
negative correlation does exist between the environmental targets set for female
productivity, and over time could be us to incrementally adjust targets if necessary.
Lastly, to improve data tracking in the program, I recommend reinstating the use of
original Egg and Larvae Data Sheets and reintroducing morphometric data collection.
83
Current data forms would need further updates to address larval count discrepancies
occurring since the application of these new data forms. Egg & Larvae Data Sheets track
important information that could be imperative to further research, including detailed
development times. These forms could be used in conjunction with current forms and
cross-referenced as needed to understand larval count discrepancies. Morphometric data
is commonly used to catch and understand divergencies between wild and captive
populations (Lewis and Thomas, 2001; Dzurisin, 2005; Crone, Pickering and Schultz,
2007). Introducing egg measurements and adult body area measurements as well are
reinstating pupal weight, adult weight, and wing measurements would help better
understand the program, and population, success at MCCCW and within other facilities.
84
CONCLUSION
This study shows that the current environmental targets for different life stages in
captivity are often not met, perhaps fortunately with no clear indication of a negative
effect on E. e. taylori survival and productivity. With the methods employed in this study,
the % of time targets are met is not an exceptional indicator of program success.
Outcomes do show an area of the program that could be expanded upon; since the
Technicians are the front line for daily care and management, providing clear absolute
targets and more focused tasks for how to achieve desired results of productivity and
survival. Continuing the research of preferred E. e. taylori environmental conditions by
looking at current field environmental conditions could change how these targets are
utilized, becoming a more reliable indication of success by making them much more
precise and accurate.
Further research into the impacts of environmental conditions on development
times and morphometric data from seasons could help illuminate the fitness. However,
the overall success of this program reveals the importance of rearing wildlife in ambient
conditions—even if there is not a complete picture of the species’ wild habitat
requirements—especially when the subjects are ectotherms that require sunlight at almost
all life stages. In addition to this captive rearing facility preventing the loss of an
endangered species and advancing E. e. taylori conservation objectives, the opportunity
for collaboration, education, and research continues to broaden the knowledge available
on E. e. taylori.
85
86
APPENDIX A – Additional Tables
Table 15. Captive rearing productivity and survival targets for seasons 2014-2015 to 2018-2019.
Seasons
2014-2015
2015-2016
2016-2017
2017-2018
2018-2019
Eggs
5000
5000
5000
5000
5000
Larvae
Retained Larvae
2500
300
2500
300
2500
300
2500
300
1800
300
Table 16. Example of a daily minimum, maximum, and average temperature and humidity table experiences duirng
captive rearing. To the right it the comparison of the absolute targets to the minimum and maximum temperature.
Date
4/25/20
4/26/20
4/27/20
4/28/20
4/29/20
4/30/20
5/1/20
5/2/20
5/3/20
5/4/20
5/5/20
5/6/20
5/7/20
5/8/20
5/9/20
5/10/20
5/11/20
5/12/20
5/13/20
5/14/20
5/15/20
5/16/20
Min
Max
Avg
Min
Max Avg
Min Max Avg
Temp Temp Temp Rh
Rh
Rh
64
72
66
52
54
52
59
71
64
41
70
55
61
78
67
54
75
63
61
72
65
53
67
61
61
77
67
53
79
67
59
73
64
62
82
72
57
76
65
68
82
73
57
71
62
75
89
83
54
69
61
71
83
78
51
74
62
69
82
75
57
77
66
56
84
76
59
81
67
60
81
72
55
83
67
58
79
69
62
88
71
54
75
64
63
94
73
55
78
68
65
83
72
44
83
64
61
80
67
69
84
78
59
84
67
59
88
78
60
93
70
45
83
69
59
91
68
46
82
71
56
104
73
34
84
64
59
76
64
70
88
78
51
69
61
34
54
52
65
104
73
75
89
83
59
80
67
57
80
70
<50º
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
0
>90º
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
FALSE
Yes
FALSE
FALSE
FALSE
Yes
Yes
Yes
FALSE
4
87
Table 17. Percent relative humidity averages of the minimum, maximum, and average percent relative
humidity for life stages pupation and males for seasons 2013-2014 to 2018-2019.
Season
2013-14
Pupation
Males
Min Max Avg Min Max Avg
41
81
62
28
71
47
2014-15
2015-16
2016-17
33
26
23
94
97
100
68
66
61
39
18
21
83
100
87
65
59
62
2017-18
2018-19
34
27
100
100
77
78
30
26
97
98
65
63
Average
31
95
69
27
89
60
Table 18. Percent relative humidity averages of the minimum, maximum, and average for life stages females
and eggs & prediapause larvae for seasons 2013-2014 to 2018-2019 and 2020-2021.
Season
2013-14
2014-15
2015-16
2016-17
2017-18
2018-19
2020-21
Average
Females &
Eggs &
Oviposition
Prediapause
Min Max Avg Min Max Avg
27
68
51
27
68
51
39
83
68
50
99
77
18
100
60
26
89
63
19
91
59
21
94
66
31
98
67
34
97
64
22
86
59
19
95
64
34
89
70
24
93
65
27
88
62
29
91
64
Table 19. Percent relative humidity averages of the minimum, maximum, and average for the life stages warm
diapause, cold diapause, and postdiapause.
Season
2013-14
2014-15
2015-16
2016-17
2017-18
2018-19
2020-21
Average
88
Warm Diapause
Cold Diapause
Postdiapause
Min Max Avg Min Max Avg Min Max Avg
28
70
49
29
90
73
36
86
64
24
100
70
41
100
92
24
94
68
23
92
61
57
99
90
26
98
68
28
97
71
42
98
86
31
93
70
26
94
61
31
100
88
22
98
68
21
87
60
37
100
86
19
100
68
43
97
80
41
99
88
46
91
67
28
91
65
40
98
86
29
94
68
Table 20. Temperature averages of the minimum, maximum, and average for the life stages males and pupation.
Season
2013-14
2014-15
2015-16
2016-17
2017-18
Males
Pupation
Min
Max
Avg
Min
Max
Avg
31
52
46
49
52
102
81
87
95
99
60
64
63
66
66
43
47
37
52
49
79
92
80
99
91
60
63
55
65
63
89
92
64
64
53
47
97
90
65
62
2018-19 49
Average 47
Table 21. Overall temperature averages of the minimum, maximum, and average temperature for life stages females &
oviposition and eggs & prediapause.
Season
2013-14
2014-15
2015-16
2016-17
2017-18
Females &
Oviposition
Min Max Avg
31
101 62
52
81
64
46
87
64
41
100 64
55
91
66
2018-19 49
2020-21 51
Average 46
95
104
94
67
67
65
Eggs & Prediapause
Min
31
53
55
48
55
Max
105
91
95
101
93
Avg
65
66
67
65
70
47
47
48
96
97
97
67
67
67
Table 22. Overall temperature averages of the minimum, maximum, and average temperature for life stages warm
diapause, cold diapause, and postdiapause.
Season
2013-14
2014-15
2015-16
2016-17
2017-18
2018-19
2020-21
Average
Warm Diapause
Min
46
51
51
52
58
51
53
52
Max
104
100
99
89
93
97
99
97
Avg
70
70
71
67
73
70
68
70
Cold Diapause
Min
13
21
22
17
17
14
26
19
Max
78
80
87
82
88
84
86
84
Avg
44
44
46
45
43
44
45
44
Postdiapause
Min
35
39
37
44
32
47
43
40
Max
79
86
85
83
91
90
82
85
Avg
57
58
56
56
58
63
56
58
89
Table 23. Number of eggs and postdiapause larvae per season. Note, prediapause release occurred for captive
populations, and 20-21 season totals include larvae in the second greenhouse at MCCCW.
Season
Wild
Captive
Total
# Eggs
# Out of # Eggs
# Out of # Eggs # Out of
Diapause
Diapause
Diapause
2013-14
2014-15
2015-16
2016-17
2017-18
2018-19
2020-21
1373
2327
2772
1261
2938
2944
2414
1324
1909
2111
1426
3016
2888
2279
2378
2437
2205
1121
609
4965
--
Table 24. Example of morphometric data summary table.
90
1456
685
624
541
146
1659
--
3751
4764
4977
2382
3547
7909
4696
2780
2594
2735
1967
3162
4547
3813
APPENDIX B – ADDITIONAL FIGURES
Figure 10. Example of MCCCW data collection form used during captive rearing.
91
Figure 11. Example snapshot of the hour environmental data analysis into the percentages of time environmental
targets were met.
92
Figure 12. Legend for the percent of time the
environmental targets are met.
Figure 13. Percent of time environmental targets were met during males for all seasons.
93
Figure 14. Percent of time females & oviposition environmental targets were met for all seasons.
Figure 15. Percent of time eggs & prediapause larvae environmental targets were met during for all seasons.
94
Figure 16. Percent of time warm diapause environmental targets were met during all seasons.
Figure 17. Percent of time cold diapause environmental targets were met for all seasons.
95
Figure 18. Percent of time postdiapause environmental targets were met for all seasons.
Figure 19. Percent of time pupation environmental targets were met for all seasons.
96
Figure 20. Legend for the Percent of days the temperature during
captive rearing is outside of the temperature targets.
Figure 21. Percent of days the minimum and maximum daily temperature is outside the male temperature target.
97
Figure 22. Percent of days the minimum and maximum daily temperature is outside the females &
oviposition temperature target.
Figure 23. Percent of days the minimum and maximum daily temperature is outside the eggs & prediapause larvae
temperature target.
98
Figure 24. Percent of days the minimum and maximum daily temperature is outside the warm diapause
temperature target.
Figure 25. Percent of days the maximum daily temperature is outside the cold diapause temperature target.
99
Figure 26. Percent of days the minimum and maximum daily temperature is outside the postdiapause
temperature target.
Figure 27. Percent of days the minimum and maximum daily temperature is outside the pupation temperature target.
100
Figure 28. Spearman's rho scatterplot matrix for percent males
productive versus percent of time environmental targets were met.
Figure 29. Spearman’s rho scatterplot matrix for percent captive females
productive versus percent of time environmental targets were met.
101
Figure 30. Spearman’s rho scatterplot matrix for the percent of captive
prediapause larvae versus percent of time environmental targets were
met.
Figure 31. Spearman's rho scatterplot matrix for the percent of captive
larvae into diapause versus percent of time environmental targets were
met.
102
Figure 32. Spearman's rho scatterplot matrix for the percent of captive
larvae out of diapause versus the percent of time environmental targets
were met.
Figure 33. Spearman's rho scatterplot matrix for the percent of captive
larvae to release versus percent of time environmental targets were met.
103
Figure 34. Spearman’s rho scatterplot matrix for the percent of wild
females productive versus percent of time environmental targets were
met.
Figure 35. Spearman's rho scatterplot matrix for the percent of the wild
prediapause larvae percent of time the environmental targets were met.
104
Figure 36. Spearman's rho scatterplot matrix for the percent of wild
larvae into diapause versus percent of time environmental targets were
met.
Figure 37. Spearman's rho scatterplot matrix for the percent of wild
larvae out of diapause versus percent of time environmental targets were
met.
105
Figure 38. Spearman's rho scatterplot matrix for the percent of wild
larvae to release versus the percent of time environmental targets were
met.
Figure 39. Spearman's rho scatterplot matrix for the percent of wild
larvae to pupation (minus the percent of larvae that entered 2nd
diapause) versus the percent of time environmental targets were met.
106
Figure 40. Spearman's rho scatterplot matric for the percent of wilds that
entered 2nd diapause (minus the percent of larvae that pupated) versus
the percent of time environmental targets were met.
Figure 41. Spearman's rho scatterplot matrix for the percent of pupae the
successfully eclosed versus the percent of time environmental targets
were met.
107
Figure 42. Spearman's rho scatterplot matrix of the percent of males
productive versus the percent days outside environmental targets.
Figure 43. Spearman's rho scatterplot matrix of the percent of captive
females productive versus the percent days outside environmental
targets.
108
Figure 44. Spearman's rho scatterplot matrix of the percent of captive
prediapause larvae versus the percent days outside environmental
targets.
Figure 45. Spearman's rho scatterplot matrix of the percent captive
larvae into diapause versus the percent days outside environmental
targets.
109
Figure 46. Spearman's rho scatterplot matrix of the percent of captive
larvae out of diapause versus the percent days above environmental
target.
Figure 47. Spearman's rho scatterplot matrix of the percent of captive
larvae to release versus the percent days outside environmental targets.
110
Figure 48. Spearman's rho scatterplot matrix of the percent of wild
females productive versus the percent days outside environmental
targets.
Figure 49. Spearman's rho scatterplot matrix of the percent of wild
prediapause larvae versus the percent days outside environmental
targets.
111
Figure 50. Spearman's rho scatterplot matrix of the percent of wild
larvae into diapause versus the percent days outside environmental
targets.
Figure 51. Spearman's rho scatterplot matrix of the percent of wild
larvae out of diapause versus the percent days above environmental
target.
112
Figure 52. Spearman's rho scatterplot matrix of the percent of wild larvae
released/retained versus the percent days outside environmental targets.
Figure 53. Spearman's rho scatterplot matrix of the percent of wild larvae
pupated versus the percent days outside environmental targets.
113
Figure 54. Spearman's rho scatterplot matrix of the percent of wild larvae
return to diapause versus the percent days outside environmental targets.
Figure 55. Spearman's rho scatterplot matrix of the percent of wild pupae
successfully eclose versus the percent days outside environmental targets.
114
Figure 56. Adult female butterfly weights (captive and wild) in the 2013-2014 season
(p=0.03, Wilcoxon rank sum test). The horizontal line inside each diamond is the group
mean.
Figure 57. Adult female butterfly weights (captive and wild) in the 2014-2015 season
(p<0.0001, Wilcoxon rank sum test. The horizontal line inside each diamond is the group
mean.
115
Figure 58. Adult female butterfly weights (captive and wild) in the 2015-2016 season
(p<0.0001, Wilcoxon rank sum test. The horizontal line inside each diamond is the
group mean.
Figure 59. Adult female butterfly weights (captive and wild) in the 2016-2017 season
(p<0.0001, Wilcoxon rank sum test. The horizontal line inside each diamond is the
group mean.
116
Figure 60. Adult female butterfly weights (captive and wild) in the 2017-2018 season
(p<0.0001, Wilcoxon rank sum test. The horizontal line inside each diamond is the
group mean.
Figure 61. Adult female butterfly weights (captive and wild) in the 2018-2019 season
(p<0.0001, Wilcoxon rank sum test. The horizontal line inside each diamond is the
group mean.
117
Figure 62. Adult female butterfly wing area (captive and wild) in the 2014-2015 season
(p=0.01, Wilcoxon rank sum test). The horizontal line inside each diamond is the group
mean.
Figure 63. Adult female butterfly wing area (captive and wild) in the 2015-2016 season
(p=0.55, Wilcoxon rank sum test). The horizontal line inside each diamond is the group
mean.
118
Figure 64. Adult female butterfly wing area (captive and wild) in the 2016-2017 season
(p=0.46, Wilcoxon rank sum test). The horizontal line inside each diamond is the
group mean.
Figure 65. Adult female butterfly wing area (captive and wild) in the 2017-2018
season (p=0.004, Wilcoxon rank sum test). The horizontal line inside each diamond is
the group mean.
119
Figure 66. Adult female butterfly wing area (captive and wild) in the 2014-2015
season (p=0.27, Wilcoxon rank sum test). The horizontal line inside each diamond is
the group mean.
Figure 67. Average temperature over 2021 in Belfair, WA.
120
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