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Identification and characterization of juvenile hormone acid methyl transferase, the ultimate enzyme in the juvenile hor...

Permanent Link: http://ufdc.ufl.edu/UFE0041645/00001

Material Information

Title: Identification and characterization of juvenile hormone acid methyl transferase, the ultimate enzyme in the juvenile hormone biosynthetic pathway of Aedes aegypti.
Physical Description: 1 online resource (133 p.)
Language: english
Creator: Van Ekert, Evelien
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2010

Subjects

Subjects / Keywords: aedes, aegypti, hormone, jhamt, juvenile
Entomology and Nematology -- Dissertations, Academic -- UF
Genre: Entomology and Nematology thesis, M.S.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Juvenile hormone acid methyl transferase (JHAMT) converts juvenile hormone acid into JH during the last step of the JH biosynthetic pathway. Since JH impacts many physiological processes in insects, disruption of its pathway can be used to control insects. Aedes aegypti vectors many arboviruses such as, dengue, chikungunya and yellow fever. Efficient control of this mosquito is the only way to eliminate these diseases. Thus, characterization and identification of key enzymes that control metamorphosis and egg development in larvae and adults of this mosquito species could save many lives. We sequenced the JHAMT gene of Ae. aegypti (jmtA), built a 3 dimensional model of the enzyme studying conformational changes as related to substrate specificity and expressed it in Escherichia coli. Following expression of JHAMT, the recombinant protein was purified by nickel affinity chromatography, stabilized in glycerol and mercaptoethanol and its specific activity was determined. The purified enzyme specificity to various substrates was tested by constructing Lineweaver Burk reciprocal plots and calculating the Km and Vmax of different substrates. JH III acid and JH I acid (cis/trans/trans isoform) were found to be excellent substrates (Vmax of 69.54 plus or minus 4.91 and 46.72 plus or minus 11.52 mmol/mol enzyme/min plus or minus SEM, respectively). Farnesoic acid and JH I acid (cis/trans/cis isoform) are good substrates (Vmax of 13.00 plus or minus 0.33 and 14.71 plus or minus 1.56 mmol/mol enzyme/min plus or minus SEM, respectively). Homo farnesoate is a moderate substrate (Vmax = 9.62 plus or minus 0.57 mmol/mol enzyme/min plus or minus SEM) whereas JH I bisepoxide acid JH III bisepoxide acid and JH I acid (trans/cis/cis isoform) are poor substrates (Vmax of 0.66 plus or minus 0.01, 0.59 plus or minus 0.07 and 0.37 plus or minus 0.03 mmol/mol enzyme/min plus or minus SEM, respectively). The activity of JHAMT was followed in vitro and in vivo using radioactively labeled 3H-methylS-adenosyl methionine (SAM) showing that the enzyme is active throughout the adult life cycle of female Ae. aegypti. RNA mediated interference (RNAi) studies using jmtA dsRNA in female Ae. aegypti inhibited egg development, whereas feeding larvae long hair pin (LHP) RNA delayed adult emergence, caused mortality and prolonged the larval stage for up to 3 weeks past normal pupation time. Following pupation, the newly emerged adults died without taking a blood meal. Tissue specific expression of jmtA was followed by Northern blot analyses during the different life stages of Ae. aegypti. This work shows that jmtA is a constitutive enzyme expressed during the different life stages of female Ae. aegypti. Although jmtA is mainly expressed in the corpora allata (CA) of female Ae. aegypti, it is also highly expressed in the ovaries of adult females, reaching a peak at 72 hours after the blood meal.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Statement of Responsibility: by Evelien Van Ekert.
Thesis: Thesis (M.S.)--University of Florida, 2010.
Local: Adviser: Borovsky, Dov.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2012-04-30

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2010
System ID: UFE0041645:00001

Permanent Link: http://ufdc.ufl.edu/UFE0041645/00001

Material Information

Title: Identification and characterization of juvenile hormone acid methyl transferase, the ultimate enzyme in the juvenile hormone biosynthetic pathway of Aedes aegypti.
Physical Description: 1 online resource (133 p.)
Language: english
Creator: Van Ekert, Evelien
Publisher: University of Florida
Place of Publication: Gainesville, Fla.
Publication Date: 2010

Subjects

Subjects / Keywords: aedes, aegypti, hormone, jhamt, juvenile
Entomology and Nematology -- Dissertations, Academic -- UF
Genre: Entomology and Nematology thesis, M.S.
bibliography   ( marcgt )
theses   ( marcgt )
government publication (state, provincial, terriorial, dependent)   ( marcgt )
born-digital   ( sobekcm )
Electronic Thesis or Dissertation

Notes

Abstract: Juvenile hormone acid methyl transferase (JHAMT) converts juvenile hormone acid into JH during the last step of the JH biosynthetic pathway. Since JH impacts many physiological processes in insects, disruption of its pathway can be used to control insects. Aedes aegypti vectors many arboviruses such as, dengue, chikungunya and yellow fever. Efficient control of this mosquito is the only way to eliminate these diseases. Thus, characterization and identification of key enzymes that control metamorphosis and egg development in larvae and adults of this mosquito species could save many lives. We sequenced the JHAMT gene of Ae. aegypti (jmtA), built a 3 dimensional model of the enzyme studying conformational changes as related to substrate specificity and expressed it in Escherichia coli. Following expression of JHAMT, the recombinant protein was purified by nickel affinity chromatography, stabilized in glycerol and mercaptoethanol and its specific activity was determined. The purified enzyme specificity to various substrates was tested by constructing Lineweaver Burk reciprocal plots and calculating the Km and Vmax of different substrates. JH III acid and JH I acid (cis/trans/trans isoform) were found to be excellent substrates (Vmax of 69.54 plus or minus 4.91 and 46.72 plus or minus 11.52 mmol/mol enzyme/min plus or minus SEM, respectively). Farnesoic acid and JH I acid (cis/trans/cis isoform) are good substrates (Vmax of 13.00 plus or minus 0.33 and 14.71 plus or minus 1.56 mmol/mol enzyme/min plus or minus SEM, respectively). Homo farnesoate is a moderate substrate (Vmax = 9.62 plus or minus 0.57 mmol/mol enzyme/min plus or minus SEM) whereas JH I bisepoxide acid JH III bisepoxide acid and JH I acid (trans/cis/cis isoform) are poor substrates (Vmax of 0.66 plus or minus 0.01, 0.59 plus or minus 0.07 and 0.37 plus or minus 0.03 mmol/mol enzyme/min plus or minus SEM, respectively). The activity of JHAMT was followed in vitro and in vivo using radioactively labeled 3H-methylS-adenosyl methionine (SAM) showing that the enzyme is active throughout the adult life cycle of female Ae. aegypti. RNA mediated interference (RNAi) studies using jmtA dsRNA in female Ae. aegypti inhibited egg development, whereas feeding larvae long hair pin (LHP) RNA delayed adult emergence, caused mortality and prolonged the larval stage for up to 3 weeks past normal pupation time. Following pupation, the newly emerged adults died without taking a blood meal. Tissue specific expression of jmtA was followed by Northern blot analyses during the different life stages of Ae. aegypti. This work shows that jmtA is a constitutive enzyme expressed during the different life stages of female Ae. aegypti. Although jmtA is mainly expressed in the corpora allata (CA) of female Ae. aegypti, it is also highly expressed in the ovaries of adult females, reaching a peak at 72 hours after the blood meal.
General Note: In the series University of Florida Digital Collections.
General Note: Includes vita.
Bibliography: Includes bibliographical references.
Source of Description: Description based on online resource; title from PDF title page.
Source of Description: This bibliographic record is available under the Creative Commons CC0 public domain dedication. The University of Florida Libraries, as creator of this bibliographic record, has waived all rights to it worldwide under copyright law, including all related and neighboring rights, to the extent allowed by law.
Statement of Responsibility: by Evelien Van Ekert.
Thesis: Thesis (M.S.)--University of Florida, 2010.
Local: Adviser: Borovsky, Dov.
Electronic Access: RESTRICTED TO UF STUDENTS, STAFF, FACULTY, AND ON-CAMPUS USE UNTIL 2012-04-30

Record Information

Source Institution: UFRGP
Rights Management: Applicable rights reserved.
Classification: lcc - LD1780 2010
System ID: UFE0041645:00001


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1 IDENTIFICATION AND CHARACTERIZATION OF JUVENILE HORMONE ACID METHYL TRANSFERASE, THE ULTIMATE ENZYME IN THE JUVENILE HORMONE BIOSYNTHETIC PATHWAY OF Aedes aegypti By EVELIEN VAN EKERT A THESIS PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF MASTER OF SCIENCE UNIVERSITY OF FLORIDA 2010

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2 2010 Evelien Van Ekert

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3 To Keith

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4 ACKNOWLEDGMENTS First of all I want to thank Professor Borovsky for giving me the opportunity to work in his laboratory. His suggestions were always very helpful and much appreciated. I want to thank professors Roug and Smagghe for their work on the molecular modeling of JHAMT, and professor Sl ma for providing us with the juvenile hormone derivatives. I also want to thank my parents to support me in my choice to work towards a degree at the University of Florida. I know it has been difficult for them to know that their little girl has moved across the ocean. But like the Puddle of Mud song states: Ther es oceans in between us, but thats not very far! I also want to thank my sisters, brother, their partners and my nephews and niece for always being there when I need them. They always make me smile and rationalize what is going on in my life. I want to thank my fianc, Keith for being patient with me. Despite having so many lonely days, nights and weekends, he has always supported me. On one of our first dates I caught a bug for my insect collection and I know he knew then I was the one. I want to th ank God for giving me the intellectual capacities to pursue an advanced education. Thank you for doing the rest (My mom always says: do your best, God will do the rest.

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5 TABLE OF CONTENTS page ACKNOWLEDGMENTS .................................................................................................. 4 LIST OF TABLES ............................................................................................................ 8 LIST OF FIGURES .......................................................................................................... 9 LIST OF ABBREVIATIONS ........................................................................................... 12 ABSTRACT ................................................................................................................... 15 CHAPTER 1 JUVENILE HORMONE ........................................................................................... 17 Introduction ............................................................................................................. 17 Juvenile Hormones ................................................................................................. 18 JH III Biosynthetic Pathway .................................................................................... 18 First Step: The Mevalonate Pathway ................................................................ 18 Second Step: Insect Specific Pathway ............................................................. 19 Juvenile Hormone Metabolism ................................................................................ 19 Juve nile Hormone Mode of Action .......................................................................... 20 Regulation of JH biosynthesis ................................................................................. 20 Juvenile Hormone Titer in Ae. aegypti .................................................................... 21 Juvenile Hormone Functions .................................................................................. 22 Develo pment .................................................................................................... 22 Female Receptivity and Reproduction .............................................................. 24 2 VECTOR CONTROL .............................................................................................. 31 Aedes aegypti ......................................................................................................... 31 Introduction ....................................................................................................... 31 Life cycle .......................................................................................................... 31 Vector Control ......................................................................................................... 32 Introduction ....................................................................................................... 32 Chemical Insecticides ....................................................................................... 32 Bio rational Insecticides .................................................................................... 33 3 MATERIALS AND METHODS ................................................................................ 35 Experimental Insects ............................................................................................... 35 RNA Isolation from Mosquitoes .............................................................................. 35 RNA Isolation from Yeast Cells ............................................................................... 35 jmtA (JHAMT gene of Ae.aegypti ) cDNA cloning .................................................... 36 Expression of jmtA in Bacterial Cells ...................................................................... 40

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6 JHAMT Expression ........................................................................................... 40 Extraction of JHAMT from Bacterial Cells ......................................................... 41 Purification of JHAMT ....................................................................................... 42 Molecular Modeling of JHAMT ................................................................................ 43 Radioactive bioassay for JHAMT ............................................................................ 45 Determination of JHAMT Activity ...................................................................... 45 Linearity of JHAMT Activity ............................................................................... 46 Effect of Metal Ions on JHAMT Activity ............................................................ 46 Substrates specificity of JHAMT ....................................................................... 47 Effect of Increasing Concentrations of S Adenosyl L methionine [methyl -3H] on JHAMT Activity ......................................................................................... 48 Effect of JH III on JHAMT ................................................................................. 48 Effect of JH III Bisepoxide Acid on JHAMT Activity .......................................... 49 In Vivo Biosynthesis of JH III from JH III acid by JHAMT ........................................ 49 Synthesis of [3H]JH III Acid ............................................................................... 49 In Vivo Biosynthesis of JH III by JHAMT in ligated females .............................. 50 In Vitro Biosynthesis of JH III by JHAMT ............................................................... 50 RNA Mediated Interference Studies ....................................................................... 51 Synthesis of jmtA dsRNAs ................................................................................ 51 Injection of dsRNA into Female Ae. aegypti ..................................................... 52 Feeding Female Mosquitoes dsRNA ................................................................ 53 RNA Mediated Interference Studies on Mosquito Larvae with Long Hairpin (LHP) RNA .................................................................................................... 54 Northern Blots Analyses of jmtA mRNA .................................................................. 55 4 RESULTS ............................................................................................................... 62 cDNA Sequence of jmtA ......................................................................................... 62 Expression of jmtA in Bacterial Cells ...................................................................... 62 Purification of JHAMT ....................................................................................... 62 SDS PAGE and Mass Spectra Analyses of JHAMT ......................................... 63 Molecular Modeling of JHAMT ................................................................................ 63 Activity of JHAMT ................................................................................................... 64 JHAMT Activity Using a Rapid Biphasic Separation (RBS) .............................. 65 Linearity of JHAMT Activity ............................................................................... 66 JHAM T Activity in the Presence of EDTA and Metal Ions ................................ 66 Substrate Specificity ......................................................................................... 67 The Effect of SAM on JHAMT Activity .............................................................. 68 Effect of JH III on JHAMT ................................................................................. 69 Effect of JH III Bisepoxide Acid on JHAMT ....................................................... 69 In Vivo Biosynthesis of JH III .................................................................................. 70 Synthesis of [3H]JH III Acid ............................................................................... 70 In Vivo Biosynthesis of [3H]JH III from [3H]JH III Acid ....................................... 70 In Vitro Biosynthesis of JH III from JH III Acid ......................................................... 71 RNA Mediated Interferenc e Studies ....................................................................... 72 Injecting dsRNA ................................................................................................ 72 Feeding dsRNA ................................................................................................ 73

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7 LHP RNA Mediated Interference of Larvae ...................................................... 74 Northern Blot Analyses ........................................................................................... 76 5 DISCUSSION ....................................................................................................... 114 LIST OF REFERENCES ............................................................................................. 126 BIOGRAPHICAL SKETCH .......................................................................................... 133

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8 LIST OF TABLES Table page 4 1 Purification of JHAMT ......................................................................................... 84 4 2 Michaelis constants (Km) and Vmax for JHAMT substrates with low concentrations of SAM ( 0.141 M) .................................................................... 97 4.3 Michaelis constants (Km) and Vmax for JHAMT substrates with saturating concentrations of SAM (600 M). ....................................................................... 97 4 4 Egg development in female Ae.aegypti injected with jmtA dsRNA. .................. 102 4 5 The effect of jmtA dsRNA and other supplements on survival and egg development in Ae. aegypti. ............................................................................. 104 4 6 Average 50% pupation time (PT) for larvae fed with a brewers yeast solution (2 % ), P. pastoris cells expressing jmtA LHP RNA and induced for 24 hours, 48 hours, 72 hours and 96 hours. ..................................................................... 110

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9 LIST OF FIGURES Figure page 1 1 Schematic representation of the neurohemal organs in the insects. .................. 27 1 2 The Juvenile Hormone group ............................................................................. 27 1 3 First step in the JH biosynthetic pathway: Conversion of acetyl CoA into Isopentenyl pyrophosphate ................................................................................ 28 1 4 Insect specific, second step in the JH biosynthetic pathway. Conversion of IPP into JH III. ..................................................................................................... 29 1 5 Juvenile hormone metabolism ............................................................................ 30 1 6 Regulation of JH biosynthesi s ............................................................................ 30 2 1 Ae. aegypti the yellow fever mosquito. .............................................................. 34 3 1 Multiple sequence alignment (MSA) of JHAMT amino acid sequences of Drosophila melanogaster Bombyx mori and Anopheles gambiae. .................... 56 3 2 Sequencing strategy of Ae. aegypti 1410 bp jmtA transcript. ............................. 56 3 3 Schematic representation of jmtA in pCR2.1. ..................................................... 57 3 4 Schematic representation of cloning jmtA in pETDuet 1 (Novagen). .................. 57 3 5 Cloning strategy of jmtA (start stop) in pET Duet 1 (Novagen). .......................... 57 3 6 JH acid derivatives used for JHAMT activity. ...................................................... 58 3 7 Chemical conversion of [3H]JH III into [3H]JH III acid by methanolic alkaline hydrolysis ........................................................................................................... 58 3 8 Synthesis of [3H]JH III by JHAMT from [3H]JH III acid. ....................................... 59 3 10 Small container with capillary tube anchored by a cotton plug (left), and female mosquitoes feeding from a capillary tube (right). .................................... 60 3 11 Long hairpin (LHP) of jmtA. ................................................................................ 60 3 12 Schematic representation of LHP cloned in pPicZB showing the cloning restriction sites ( Xho I and Xba I) the AOX1 promoter (P) and Zeocin resistant gene (Zeo). ......................................................................................................... 61 4 1 Nucleotide sequence of jmtA cDNA (Borovsky et al. 2006) (GenBank accession number DQ 409061). ......................................................................... 78

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10 4 2 Genomic DNA sequence of jmtA including introns (black), termination region and TATA and CAAT boxes on the promoter region. ......................................... 79 4 3 Nickel column purification of JHAMT. ................................................................. 79 4 4 SDS polyacrylamide gel electrophoresis of fractions obtained by nickel column purification. ............................................................................................. 80 4 5 Mass spectrometry of the 32 kDa band excised from the SDS gel. .................... 80 4 6 Molecular modeling of Ae. aegypti JHAMT. ........................................................ 81 4 7 C18 RP HPLC of JHIII and MF standards. .......................................................... 83 4 8 Conversion of FA into MF by purified JHAMT (blue line) and crude extract (red line). ............................................................................................................ 83 4 9 Synthesis of [3H]JH III and [3H]MF from JH III acid and FA respectively with purified JHAMT (4 g) and 1.1 Ci (0.141 M) [3H]SAM. .................................... 84 4 10 Linear relationship between JHAMT activity and time. ....................................... 85 4 11 Activity of JHAMT in the presence and absence of EDTA. ................................. 86 4 12 Activity of JHAMT in different salt solutions. ....................................................... 87 4 13 Substrate specificity of Ae.aegypti JHAMT in the presence of SAM. .................. 88 4 14 Lineweaver Burk reciprocal plots using low SAM concentrations (0.141 M) ..... 89 4 15 Lineweaver Burk reciprocal plots using saturating SAM concentrations (600 M) ..................................................................................................................... 93 4 16 Activity of JHAMT with increasing concentrations of JH III acid .......................... 98 4 17 Synthesis of MF from FA by JHAMT in the presence of increasing concentrations of JH III (0 3.75 mM). ................................................................. 99 4 18 Synthesis of JH III from JH III acid by JHAMT in the presence of increasing concentrations of JH III (0 3.75 mM). ................................................................. 99 4 19 Biosynthesis of JH III and MF from JH III acid and FA in the presence of JH III (1375 M) (blue line) and in the absence of JH III (red line). ........................ 100 4 20 Synthesis of [3H] JH III by JHAMT in the presence of increasing concentrations of JH III bisepoxide acid (02000 M). ...................................... 100 4 21 Conversion of [3H] JH III into [3H]JH III acid by alkaline hydrolysis. .................. 101

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11 4 22 In vivo conversion of [3H]JH III acid into [3H]JH III by JHAMT. ......................... 101 4 23 In vitro biosynthesis of JH III by extracts of Ae. aegypti headthoraces (HT) and ovaries (OV). ............................................................................................. 102 4 24 Light microscopy of yolk development in ovaries of jmtA dsRNA and water injected Ae. aegypti ......................................................................................... 103 4 25 Average yolk length of jmtA dsRNA injected and water injected (control) female Ae. aegypti ........................................................................................... 103 4 26 Light microscopy of undev eloped ovaries removed from female Ae. aegypti 72 hours after the blood meal that was fed jmtA dsRNA for 7 days before the blood meal. ....................................................................................................... 104 4 27 Effect of jmtA dsRNA on egg development after a second blood meal. ........... 105 4 28 Ovaries and oocytes f rom mosquitoes fed jmtA dsRNA during the second gonodotrophic cycle. ......................................................................................... 106 4 29 Pupation periods for larvae ............................................................................... 108 4 30 Survival of larvae fed P. pastoris cells transformed with jmtA LHP RNA and with non transformed yeast cells. ..................................................................... 110 4 31 Survival of larvae that were fed jmtA LHP RNA yeast cells. ............................. 111 4 32 Northern blot analyses of head thoraces and abdomens during different stages of the gonadotrophic cycle of female Ae. aegypti ................................. 112 4 33 Northern blot analyses of ovaries 3 days post emergence (PE), 48 h post the blood meal (PBM) and 72 h PBM. .................................................................... 112 4 34 Northern blot analyses of ovaries (OV) 72 h PBM from mosquitoes fed with 5% sugar solution (control) and mosquitoes fed with 60 g/10 l dsRNA. ....... 113 5 1 View of JHAMT molecule showing JH III Acid binding cavity, SAM binding site and a secondary groove that allow JH III acid molecules to bind non speci fically. ....................................................................................................... 125 5 2 Last steps of the JH III biosynthetic pathway. ................................................... 125

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12 LIST OF ABBREVIATION S ACN acetonitrile Ae. aegypti Aedes aegypti A. gambiae Anopheles gambiae BME mercaptoethanol BMGY buffered minimal growth medium for yeast BMM buffered minimal medium B. mori Bombyx mori B.t. Bacillus thuringi ensis B.t.i. Bacillus thuringi ensis variety israelensis CA corpus allatum (singular), corpora allata (plural) Cb SAMT S Adenosyl methioninedependent methyl transferase from Clarkia breweri CC cor pus cardiacum (singular), corpora cardiaca (plural) cDNA complementary DNA DDT dichl orodiphenyltrichloroethane DEPC diethyl pyrocarbonate DMAP dimethylallyl diphosphate DMSO dimethyl sulfoxide D. melanogaster Drosophila melanogaster DTT d ithiothreitol E. coli Escherichia coli EDTA ethylenediaminetetraacetic acid EST expressed sequence tag FA farnesoic acid FPP farnesyl diphosphate

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13 HCA h ydrophobic cluster a nalysis HPLC h igh performance liquid chromatography IGR insect growth regulator IPP isopentenyl diphosphate IPTG i sopropyl -D1 thiogalactopyranoside JH juvenile hormone JHA juvenile hormone acid JHAMT juvenile hormone acid methyl transferase JHBP juvenile hormone binding protein JHE juvenile hormone esterase JHEH juvenile hormone epoxide hydr olase jmtA JHAMT gene of Ae.aegypti LB lysogeny broth LHP long hairpin MF methyl farnesoate MGY minimal growth medium for yeast MM minimal medium MMLV Molony murine leukemia virus NSC neurosecretory cell PCR polymerase chain reaction PG prothoracic gland PI pars intercerbralis PMSF phenylmethanesulphonylfluoride P. pastoris Pichia pastoris 50 PT 50% pupation time

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14 PTTH prothoracicotrophic hormone PV parental vector (pPicZB without insert) RACE rapid amplification of cDNA ends RBS rapid biphasic separation RNAi RNA mediated interference SAM S adenosyl methionine S AM Mts S adenosylmethioninedependent methyltransferases. SDS PAGE sodium dodecyl sulfate polyacrylamide gel electrophoresis TIGR the institute of genomic research UTR untranslated region

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15 Abstract of Thesis Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Master of Science IDENTIFICATION AND CHARACTERIZATION OF JUVENILE HORMONE ACID METHYL TRANSFERASE, THE ULTIMATE ENZYME IN THE JUVENILE HORMONE BIOSYNTHETIC PATHWAY OF A edes aegypti By Evelien Van Ekert May 2010 Chair: Dov Borovsky Major: e ntomology and nematology Juvenile hormone a cid m ethyl t ransferase (JHAMT) converts juvenile hormone acid into JH during the last step of the JH biosynthetic pathway. Since JH i m pact s many physiological processes in insects, disruption of its pathway can be used to control insects. A e des aegypti vectors many arboviruses such as, dengue, chikungunya and yellow fever. Efficient c ontrol of this mosquito is the only way to eliminate the se diseases. Thus, characterization and identification of key enzyme s that control metamorphosis and egg development in larvae and adults of this mosquito species could save many lives We sequenced the JHAMT gene of Ae. aegypti ( jmtA ) built a 3 dimensional model of the enzyme studying conformational changes as related to substrate spec ificity and expressed it in Escherichia coli Following expression of JHAMT, the recombinant protein was purified by nickel affinity chromatography, stabilized in glycerol and mercaptoethanol and its specific activity was determined. The purified enzyme sp ecificity to various substrates was tested by constructing Lineweaver Burk recip r ocal plots and calculating the Km and Vmax of different substrates JH III acid and JH I acid (cis/trans/trans isoform) were found to be excellent substrates ( Vmax of

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16 69.54 4.91 and 46.72 11.52 mmol/ mol enzyme/min SEM respectively). Farnesoic acid and JH I acid (cis/trans/cis isoform) are good substrates ( Vmax of 13.00 0.33 and 14.71 1.56 mmol/mol enzyme/min SEM respectively ) Homo farnesoate is a moderate substr ate ( Vmax = 9.62 0.57 mmol/mol enzyme/min SEM ) whereas JH I bisepoxide acid JH III bisepoxide acid and JH I acid (trans/cis/cis isoform) are poor substrates ( Vmax of 0.66 0.01, 0.59 0.07 and 0.37 0.03 mmol/mol enzyme/min SEM respectively) The activity of JHAMT was followed in vitro and in vivo using radioactively labeled [3H methyl ] S a denosyl m eth ionine (SAM ) showing that the enzyme is active throughout the adult life cycle of female Ae. aegypt i RNA mediated interference (RNAi) studies using jmtA dsRNA in female Ae. aegypti inhibited egg development whereas feeding larvae long hair pin (LHP) RNA delayed adult emergence, caused mortality and prolonged the larval stage for up to 3 weeks past normal pupation time. Following pupation, the newly emerged adults died without taking a blood meal Tissue specific expression of jmtA was followed by Northern blot analyses during the different life stages of Ae. aegypti This work shows that jmtA is a constitutive enzyme expressed during the different life stages of female Ae. aegypti Although jmtA is mainly expressed in the corpora allata ( CA ) of female Ae. aegypti it is also highly expressed in the ovaries of adult female s, reaching a peak at 72 hours after the blood meal.

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17 CHAPTER 1 JUVENILE HORMONE Introduction Hormones are chemical messengers produced by multicellular organisms to relay to confirm the presence of insect hormones. W orking on the larvae of the gypsy moth observed that the insect s brain control s insect molting by releasing a hormone into the hemolymph. Later it was confirmed by Wigglesworth that the neurosecretory cells (NSCs) in the insect brain produce a prothoracicotropic hormone (PTTH). PTTH is a peptide hormone that is released by neurohemal organs located just behind the brain on the aorta wall and are named corpora cardiaca (CC) ( Fig. 1 1), In Manduca sexta PTTH is released by endocrine glands situated just behind the CC and are called the corpora allata (CA) PTTH binds to a receptor on the prothoracic glands (PG) and stimulates the PG to synthesi ze and release ecdysteroids. Ecdysone and its active hydroxylated derivative, 20hydroxyecdysone, have multiple effects on ecdysis and metamorphosis in a rthropods Insects do not synthesize steroids they obtain them from plants These moieties are converted into ecdysone and 20hydroxyecdysone through a series of enzymatic processes (Thummel et al. 2002) Twenty hydroxyecdysone, formed by hydroxylation of ecdysone, is mostly invol ved in inducing a molt allowing insects to shed their cuticle and grow Ecdysteroid is primarily synthesized by the PG, however, synthesis also occurs in cells of the fat body, the testes and the ovaries. The PG undergo apoptosis after metamorphosis into t he adult (Klowden 2002) ; this apoptosis is is controlled by JH. The hormone is not just responsible for the apoptosis of these glands but it is involved in many physiological processes which will be discussed in

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18 detail in the following chapters. The JHs ( JH I, II, III and 0) are sesquiterpenoids which are synthesized and released primarily by the CA In mosquitoes however, Borovsky and colleagues reported that JH III was also synthesized by the ovar ies and male accessory glands of mosquitoes (Borovsky et al. 1994a 1994b) The CA are endocrine glands located in mosquitoes behind the brain in the junction between the head and the thorax ( Fig. 1 1) (Klowden 2002) Juvenile Hormones The JHs are a group of acyclic sesquiterpenoids that are oxidized and r earranged terpenes. Sesquiterpenes are made out of three isoprene units with a molecular formula of C15H24. Th e JH group contains six major members : JH 0, JH I, JH II, JH III, 4 methyl JH I and JH III bisepoxide. JH III is the least substituted form of the JH group ( Fig. 1 2). JH III is found in most insect orders, while the homolog ues are found only in higher insect orders. In Diptera, JH III bisepoxide has been shown to be predominant however, its role is not known (Richard et al. 1989) JH I II Biosynthetic Pathway F irst S tep: T he M evalonate P athway JH III is found in all insect orders including mosquitoes. Its biosynthetic pathway starts after isopentenyl diphosphate (IPP) is produced by the mevalonate pathway; a common pathway used by bacter ia, archaea, fungi, plants and animals to synthesize terpenoids. The mevalonate pathway uses acetate as the first building block and converts it into acetyl CoA, which is then converted into acetoacetyl CoA by Claisen condensation. Addition of another acetyl CoA through an aldol like condensation and reduction forms mevalonate which is converted into IPP by phosphorylation and decarboxylation ( Fig. 1 3) (McMurry et al. 2007 Klowden 2002, Goodman et al. 2005).

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19 S econd Step: Insect Specific Pathway Following phosphorylation and decarboxylation, IPP is converted into farnesyl diphosphate (FPP) by covalently linking two IPPs and dimethylallyl diphosphate (DMAP). At this point, the biosynthetic pathway of JH is insect specific and therefore is important for future insect control. FPP is dephosphorylated forming farnesol which is dehydrogenated and converted into the aldehyde form, farnesal. Further reduction of farnesal by dehydrogenation forms farnesoic acid (FA) which can be converted into JH III by two differe nt pathways: a. FA can be first epoxidized into JH III acid and then methylated into JH III by S adenosyl methionine (SAM) dependent methyl transferase or, b. FA can be first converted by SAM mediated methyl transferase into methyl farnesoate (MF) and then epoxidized into JH III ( Fig. 1 4) (Goodman et al. 2005). Juvenile Hormone Metabolism JH titer in the hemolymph is regulated by metabolism ( Fig. 1 5 ). After JH biosynthesis decreases before metamorphosis, excess hormone will be rapidly metabolized to reduc e its level in the hemolymph. JH metabolism occurs in the hemolymph or in insect cells after they have sequestered the hormone. JH is inactivated by JH esterase (JHE) and JH epoxide hydrolase (JHEH) Two pathways have been suggested for JH metabolism: a. JH is first hydrolyzed into JH acid by JHE and then converted into JH acid diol by JHEH and, b. JH epoxide ring is first rehydrated and converted into a diol by JHE H followed by hydrolysis of the methyl ester group by JHE into JH acid diol (De Kort et al. 1981) In A e aegypti JH is first metabolized by JHE into JH acid before it is metabolized by JHEH into the diol acid (Borovsky et al. 1992) whereas in Culex quinquefasciatus it was suggested that JH is first converted into JH diol by JHEH and then hydrolyzed into the diol acid by JHE (Lassiter et al. 1995)

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20 Juvenile Hormone Mode of Action Despite its importance for many physiological processes, the mode of action of JH is still unclear and to date, no JH receptor has been found (Konopova et al. 2008; Wil lis 2007). After its release from the CA, JH binds to a JH binding protein (JHBP) which circulates in the hemolymph. This complex is needed to transport the hormone because JH is very lipophilic, and the binding to JHBP protects the hormone from degradatio n. Different JH binding proteins and lipophorins have been found in different insect species. The hormone protein complex targets different tissues in different insects. In Drosophila melanogaster intracellular binding sites for JH are found in the fat bo dy, epidermis, ovary and male accessory glands. In certain Lepidoptera, JH induces the transcription of the vitellogenin mRNA in the fat body (Riddiford 1994). Regulation of JH biosynthesis Because of the importance of JH in different physiological aspect s of insects, the titer of JH in the hemolymph is closely regulated either by biosynthesis or by degradation. Biosynthesis seems to be the main regulator y mechanism that maintains a certain level of JH in most insect species In Lepidoptera however, it has been shown that catabolic activity also play s a major role in maintaining JH titers Therefore, the regulation of JH metabolic enzymes plays a very important role in maintaining JH titers in insects hemolymph. Because JH is synthesized and rapidly rele ased by the CA, regulation of the CA determines the JH titer in the hemolymph of insects The CA is regulated by nerves that originate at the NSCs of the brain or by humorous factors that circulate in the hemolymph (P ric Mataruga et al. 2006) Allatotropins are neurosecretory peptides that stimulate the CA to synthesize JH and are released by the pars intercerbralis (PI) of the brain. The PI cells respond to internal and external stimuli

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21 like f eeding, mating and photoperiod. Allatoinhibins an d allatostatins are also neurosecretory peptides that are important in the regulation of the CA. The allatoinhibins, found in Lepidoptera, inhibit CA activity by making the CA irreversibly unresponsive to allatotropins in the hemolymph. Allatostatins, on t he other hand, reversibly inhibit JH biosynthesis by the CA (Stay et al. 2007). Internal mechanisms of CA regulation include negative and positive feedback by JH. When JH titer increases in the hemolymph, its biosynthesis by the CA decreases. This phenomenon was tested by Tobe and Stay by treating gonadotrophic Diploptera punctata (Dictyoptera) with a JH analogue (Tobe et al. 197 9 ) Positive feedback by JH was shown by allatectomizing half the CA. The single CA gland that was left compensated for the allate ctomized half (Stay et al. 1978) Ovaries in adult insects have also been shown to influence JH biosynthesis. Depending on the stage of the oocyte, developing ovaries can stimulate and inhibit JH biosynthesis. Similar phenomena have also been reported in D punctata (Rankin et al. 1984). JH binding proteins are also involved in the regulation of JH because they protect JH from nonspecific esterases that otherwise would break it down (Tobe et al. 1985) It appears that JH biosynthesis and its titer in the hemolymph is controlled by multiple factors and stimuli ( Fig. 1 6). Juvenile Hormone Titer in Ae. aegypti JH titer in Ae. aegypti was extensively studied by several research groups (Borovsky et al 1985 Shapiro et al 1986 Readio et al. 1988, Borovsky et al 1992, Li et al. 2003a ). The CA were shown to synthesize JH during the life cycle of adult female mosquitoes. JH titer in Ae. aegypti is high in sugar fed females and the titer drops to a low level 12 to 24 hours after the blood meal. At 3648 hours af ter the blood meal the titer increases again (Readio et al 1988). JH biosynthesis was also followed using [12-

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22 3H] MF in vivo by topical applications of MF and in vitro by incubating exposed CA with the radioactively labeled MF. In vivo studies showed that after eclosion JH III titer increased, reaching a m a ximum of 22 fmol/hour /female at 6 days. In vitro the JH titer after eclosion first dropped and then increased again until a level of 12 fmol/hour /CA was reached at 4 to 6 days after emergence. Both in vivo and in vitro results show a sharp increase in JH titer soon after the blood meal which could be due to upregulation by the blood meal of MF epoxidase. The in vivo and in vitro studies show that 4 to 10 hours after the blood meal a decline in JH I II biosynthesis occurs reaching a minimum at 24 h. The synthesis increases afterwards to prepare the ovaries and fat body for a second gonadotrophic cycle (Borovsky et al. 1992). Juvenile Hormone Functions JH is involved in many physiological processes in insects. JH influences embryonic, larval, pupal and adult development, caste determination, larval feeding rate, wandering behavior, diapause, pheromone production, female receptivity and reproduction (Jones 1995). Development Development of the insect e mbryo has been found to be regulated by JH. Newly laid eggs have high JH esterase activity which destroys maternal JH. Riddiford & Williams found that JH analogues inhibit embryonic development (Riddiford & Williams 1967) Injection of adults before ovipos ition or mating stops egg development at the blastoderm stage, and no hatching was observed (Riddiford 1994). Topical application of silkworm ( Bombyx mori Lepidoptera) eggs with JH analogues also stopped egg hatch ing (Riddiford et al. 1967) I n cat fleas ( Ctenocephalides felis, Siphonaptera) JH has little effect on egg hatching (Meola et al. 2001) whereas in the firebug ( Pyrrhocoris

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23 apterus Heteroptera) and the large milkweed bug ( Oncopeltus fasciatus Heteroptera) JH treatment of eggs caused an arrest of metamorphosis in post embryonic insects (Riddiford 1970) JH esterase activity goes down at the last third of the embryonic development and JH has been shown to be present in insect embryos. The hormone is important in cuticle formation and gut development in first instar larvae (Riddiford 1994) The development of post embryonic insects is a complex process encompassing many steps that are controlled by 20hydroxyecdysone and JH. The combination of JH and ecdysone influences the cellular comm itment of pupation and adult development, whereas the presence of JH alone prevent s melanin formation in the cuticle. Different responses have been observed in different insect orders. The hemolymph titer of JH controls the specific developmental stage of insect s. Wigglesworth showed that the removal of the CA from juvenile triatomine bug ( Rhodnius prolixus Hemiptera) caused the insect to develop prematurely into an adult at the next molt (Wigglesworth 1970) Adding JH to a last instar larva causes the ins ect to remain juvenile at the next molt and causes supernumerary instars. This has also been observed in Lepidoptera and Coleoptera (Zhou et al. 2002) In these insect orders the cuticles of the larvae, pupae and adults are made up from epidermal cells. I n D melanogaster (Diptera) addition of JH does not prevent pupation, though it does disrupt adult development of the abdominal cuticle (Postlethwait 1974) The reason for this is that the pupal epidermis, except for the abdomen, is derived from imaginal discs, and exogenous JH does not prevent the larval pupal transformation, even when applied throughout larval life. JH does not have any effect on the subsequent external adult differentiation of the head and thorax, although JH disrupts metamorphosis of the nervous and muscular systems

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24 when given during the prepupal period. However, JH application during the final larval instar or during the prepupal period prevents normal adult differentiation of the abdomen, whose cells arise from proliferation of the histoblasts during the prepupal period (Zhou et al. 2002 ) Metamorphosis takes place when there is a peak in ecdysteroids and absence of JH in the last instar larva (Klowden 2002) Female R eceptivity and R eproduction Sexual maturation is a phenomenon common to the animal world. Insects have to mature sexually before they are able to reproduce. This maturation includes the acquisition of male courtship behavior and female receptivity. Female receptivity goes hand in hand with sequestering yolk proteins (vitellogenins) by the developing oocytes. In D melanogaster JH is known to influence the synthesis of vitellogenins and switch on female receptivity (Ringo et a l. 1991, Manning 1967 ) In many insects JH has been found to be necessary for egg maturation. Egg development and maturation is a multi step process. The ovaries are made up of ovarioles which consist of a germarium and a primary follicle. This primary follicle contains one oocyte and seven nurse cells and these are surrounded by a layer of follicle cells and epithelial cells (Clements 2000 Riehle et al. 2002) Immediately after adult eclosion, the primary follicles start growing in size and number and differentiate (Hagedorn et al. 1977) JH is required to get the follicles to this previtellogenic resting stage (Masler et al. 1979) At the same time, the ovaries and fat bodies become competent for respectively uptake of vitellogenins and vitellogenin synthesis (Hagedorn, 1974, Hagedorn et al. 1977, Flanagan et al. 1977, Riehle et al. 2002) In Diptera, JH and the ecdysteroids play different roles in the egg maturation in the first gonadotrophic cycle (Qin et al. 1995) In many Diptera JH and ecdysteroids are both involved in

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25 vitellogenin bio synthesis by the fat body and vitellogenin uptake by the developing oocytes, which deposit the egg yolk protein as vitellins. In Ae aegypti the role of JH in vitellogenesis has bee n controversial (Gwadz et al. 1973, Borovsky 1984) Microsurgical experiments by Arden Lea & Meola showed that the CA are not necessary for egg development after the blood meal (Lea 1969 Meola et al. 1972). Allatectomy within 1 hour after adult emergence prevented yolk deposition in the ovary after blood feeding but allatectomy 3 days after adult emergence produced a normal clutch of eggs (Lea 1963). Since feeding nonphysiological concentrations of 20OH ecdysone stimulated yolk deposition (Spielman et a l. 1971), the question was raised if 20OH ecdysone stimulates vitellogenin synthesis in vitro and in vivo Schlaeger and collea gues found that the ecdysteroid concentrations in female Ae. aegypti increased after a blood meal (Schlaeger et al. 1974). Using tissue culture assays Hagedorn and collegues proposed that after the blood meal a brain factor, ovarian ecdysteroidogenic hormone (OEH) was released that stimulated the ovaries to secrete ecdysone in the haemolymph. The ecdysone was then converted in the fat bodies into the active 20OH ecdysone. The latter stimulated the fat bodies to synthesize and secrete vitellogenin in the haemolymph which was then sequestered by the ovaries in order to mature the eggs (Hagedorn et al. 1975). Thus, it was generally as sumed that 20hydroxyecdysone was the key factor in vitellogenesis. The model that Hagedorn proposed had several weak points. Injections of non physiological concentrations of 20OH ecdysone were needed to stimulate vitellogenesis and ovarian development, in vivo (Lea 1982). Also, fat bodies from sugar fed females could not be stimulated to synthesize vitellogenin after implantation of ovaries from a blood fed female which synthesized ecdysone (Borovsky et al. 1979).

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26 Lea and Racioppi proposed that there could be another factor synergizing with 20hydroxyecdysone that stimulates vitellogenesis (Lea 1982, Racioppi et al. 1983). Borovsky et al. (1985) proposed a synergistic effect of juvenile hormone and 20hydroxyecdysone for successful synthesis of vitellogenin. Ligated abdomens that were treated with methoprene (JH analogue) and implanted with an ovary from a blood fed donor secreting ecdysone indicated that both hormones were needed for normal egg development (Borovsky 1981). Indeed, there is an increase in JH III synthesis immediately after the blood meal (Borovsky et al. 1992). It was proposed that JH III plays a regulatory role in the stimulation of the vitellogenin gene, while 20hydroxyecdysone may have a role in the translation of the gene (Borovsky, 1985).

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27 F igure 11. Schematic representation of the neurohemal organs in the insects. LNC: lateral neurosecretory cells (NC), MNC: medial NC, ANC: anterior NC, INC: intermediate NC, VNC: ventral NC, CC: corpora cardiaca, CA: corpora allata, PG: prothoracicotrophic glands (modified from Clements 2000) Figure 12. The Juvenile Hormone group

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28 Figure 13. First step in the JH b iosynthetic pathway : Conversion of acetyl CoA into Isopentenyl pyrophosphate

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29 Figure 14. Insect specific, s econd step in the JH biosynthetic pathway C onversion of IPP into JH III.

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30 Figure 15 J uvenile hormone metabolism Figure 16 Regulation of JH biosynthesis (modified from Klowden 2002)

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31 CHAPTER 2 VECTOR CONTROL Aedes aegypti Introduction Ae. aegypti the yellow fever mosquito vectors dengue fever, chikungunya and yellow fever virus es in the tropics and subtropics. The mosquito is easily recogniz ed by its silver white scales forming a lyreshape on the scutum and its bands of white scales on each tarsal segment of the hind legs ( Fig. 2 1) (Darsie et al. 2004) The complete genome of Ae aegypti has been sequenced and was published in 2007 by scientists at the J Craig Venter Institute and the University of Notre Dame ( GenBank accession number AAGE00000000) Life cycle Female Ae. aegypti oviposit on damp surfaces in tires, jars, tree holes, cans and urns. These oviposition sites provide an excellent larval habitat and adult resting site. The eggs can remain dormant for long periods (over a year), and they hatch when submerged under water causing oxygen deprivation to the first instar larva that breaks the egg shell and emerges into the water (Gillet et al. 1977). Ae. aegypti larvae go through four larval instar stages, and when temperature and nutrition are optimal larvae become pupae in five days (Christophers 1960). Larvae die at temperat ures below 10 C and above 44 C. They feed on the microbiota and decaying material found in their habitat. When food availability is scarce during larval development, the adults exhibit low weight, decrease in longevity, sex ratio that favors males (Haramis 1983) and host seeking behav ior of starved females is less successful (Klowden et al. 1988).

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32 Vector Control Introduction Dengue fever is a painful arboviral disease occurring in the tropics and subtropics at high frequency More than 50100 million cases of dengue fever have been rep orted worldwide each year. No vaccine has been developed so far for the four distinct, but closely related arboviruses that cause the disease. Therefore, effective mosquito control is the only approach to keep the disease under control (Devine et al. 2009; WHO fact sheet) Chemical Insecticides The use of chemical insecticides dates back to the 19th century when Paris green was used against the Colorado potato beetle (McWilliams 2008) Since then, a multitude of chemical insecticides have been developed an d used against a wide variety of pests and disease transmitting insects. Ninety nine percent of the insecticides that are currently sold are chemical insecticides ( Rosell et al. 2008) The most widely known chemical insecticide against mosquitoes is dichlo rodiphenyltrichloroethane (DDT) (McWilliams 2008) This chlorinated hydrocarbon was widely used during the late 1940s and subsequently was banned by the US government in 1972 because of its potential harm to people and the environment. When c hlorinated hy drocarbons degrade in the environment their residues incorporate into the food chain and cause irreversible ecological damage Therefore other chemical insecticides were developed. These new insecticides break down faster into nontoxic moieties Organoph osphates (e.g. malathion) and carbamates (e.g. sevin) are examples of these new insecticides The organophosphates attack the nervous system of insects, while the carbamates are toxic to the different life stages of insects upon contact. Despite enormous i mprovements in

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33 the development of chemical insecticides there is still a concern that these chemicals are a potential hazard to the environment and to other organisms especially beneficial insects. R esistance against these new insecticides however, has developed by insects because of extensive and repeated use (McWilliams 2008) Bio rational Insecticides Because chemical insecticides cause health and ecological problems there is a n urgent need to develop selective insecticides with minimal environment al impact The new bio rational insecticides that are currently used are the insect growth regulators (IGR's) and Bacillus thuringiensis ( B.t. ) toxins (Factsheets: biorationals) IGRs include JH mimics and inhibitors. Because JH is an important hormone in i nsect developmental processes, the application of mimics or inhibitors causes detrimental effects on the molting and metamorphosis of insect s (Factsheets: insect products) B.t. is a group of bacteria that produce toxic proteins (Cry and Cyt). The variety israelensis ( B.t.i.) is very effective and specific against mosquito larvae. The Cry and Cyt proteins form pores in the midgut s of larvae causing loss of osmotic pressure, movement of water into the gut and rapid death (Factsheets: biopesticides) B io rati onal insecticides comprise only 1% of the insecticid al market because they act slow er as compared with organic insecticides are more expensive to produce and have limited niche applications. More research and more products are needed to develop new biora tional insecticides to replace the environmental toxic insecticides currently in use (Rosell et al. 2008)

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34 F igure 21. Ae. aegypti the yellow fever mosquito. The red circle marks the silver white scales that form a lyreshape on the scutum. The green circle marks a band of white scales on a tarsal segment of the hind legs.

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35 CHAPTER 3 MATERIALS AND METHOD S Experimental I nsects Ae. aegypti larvae were reared at 27C with a light:dark cycle of 12:12 on brewers yeast and lactalbumin ( 1:1 ; w:w) Newly emerged adults were maintained on sucrose solution (5%) adsorbed on to cotton wool pad s. Female mosquitoes were maintained on water 24 h prior to blood feeding on a chicken. RNA Isolation from Mosquitoes Mosquitoes and surgically removed mosquito tissues were homogenized in 500 L Trizol reagent (Invitrogen, CA ) and the homogenates were centrifuged at 14,000 rpm for 10 min in an Eppendorf centrifuge, at room temperature. After centrifugation, supernatants were removed and c hloroform (200 L) was added to separate RNA from DNA and proteins. The aqueous layer containing RNA was removed and the RNA precipitated with isopropyl alcohol ( 500 L ) and washed with ethanol (75% ). The RNA pellet was dissolved in deionized, diethylpyrocarbonate (DEPC) ( 2 % ) treated water and its purity and concentration determined by reading the absorbance at 260 and 280nm using a DNA quant machine ( Biochrom Ltd., Cambridge, England) RNA Isolation from Yeast Cells Yeast cells ( 1 mL ; OD600 = 40; 3 x 109 cells/ m L ) w ere centrifuged down at 14,000 rpm in an Eppendorf centrifuge. The pellet was resuspended in Trizol reagent ( 500 L ) (Invitrogen, CA ) and acid washed glass beads ( 180 m ) ( Sigma Aldrich St. Louis, MO ) were added The cells were broken in a FastPrep instrument ( Savant Instruments, Inc., Holbrook, NY ) for 40 seconds and cell walls were pelleted by centrifug ation for 10 min

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36 at 14,000 rpm in an Eppendorf centrifuge at room te mperature The supernatant was removed and Trizol reagent ( 500 L) (Invitrogen) was added to the cell wall pellet and the pellet and unbroken cells were broken again as above. The supernatant s after the 2 cellbreaking cycles were combined, and RNA was iso lated as described above (see RNA isolation from mosquitoes) jmtA (JHAMT gene of Ae.aegypti ) cDNA cloning Using multiple sequence alignment (MSA) with C lustal W, homolog ous regions were detected in JHAMT sequences of D melanogaster B. mori and Anopheles gambiae (Accession numbers AE014134.5, AB113578.1 and AAAB01008900.1, respectively) ( Fig. 3 1) Degenerate primers were designed to anneal to the homologous regions. (DB 913: 5 AAY AAR GCI AAY YTI TAY CAR 3 ( tm 36 48.9 C) ; DB 914: 5 GGI GTI CA R MGI MGI GAY GC 3 ( tm 45.9 56.2 C) ; DB915: 5 GAY ATH WSI GAR CAR ATG GT 3 ( tm 42.5 51.1 C) ; DB916: 5 TTY TAY TGY YTI CAY TGG GTI CA 3 ( tm 44.6 54.4 C) ; DB917: 5 YTG IAC CCA RTG IAR RCA RTA RAA 3 ( tm 45.5 56.7 C) ; DB918: 5 IGC RTA IAC IAC IAC IAR YTT RTA 3 ( tm 40.3 48.9 C) ( R = A/G, Y = T/C, W = T/A, H = T/C/A, M = C/A) ( Fig. 3 2) A RT PCR reaction (20 L ) containing 4 L 25 mM MgCl2, 2 L 10 x PCR buffer (Applied Biosystems, CA), 6 L sterile distilled water, 4 L dNTP mix (10 mM each of dATP, dTTP, dCTP, and dGTP), 1 L RNase inhibitor (20 U), 1 L Moloney murine leukemia virus ( MMLV ) reverse transcriptase (50 U) was prepared for each reaction containing 1 L reverse primer (DB917, DB 918) ( 15 M ) and total RNA (1 g) R everse transcription (RT) was per formed in a DNA thermal cycler (Applied Biosystems, CA USA ) at 24 C for 10 min, followed by 42 C for 60 min, 52 C for 30 min, 99 C for 5 min, and 5 C for 5 min After RT, 3 L 10 x Buffer, 25.5 L sterile distilled water, 0.5 L

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37 AmpliTaq DNA polymerase (2.5 U) and 1 L (15 M) of the forward primers DB 913, DB914, DB915, DB916 were added to each reaction. PCR was carried out as follows: denaturation for 3 min at 95 C (1 cycle), annealing for 4 min at 48 C extension for 40 min at 60 C (1 cycle each), denaturation at 95 C for 30 seconds annealing for 30 seconds at 48 C and extension for 2 minutes at 60 C (40 cycles) with a final extension for 15 min at 60 C After RT PCR, the amplified ds DNA in each reaction tube was stored at 20 C until use. A small sequence of the jmtA gene of Ae. aegypti was discovered in expressed sequence tag ( EST ) libraries of the institute of genomic research ( TIGR ) by blasting the library with the D. melanogaster seq uence. Genespecific primers were synthesized (DB932: 5 GCG TTT TCC AAC ATT TAT AAT CTT 3 ( tm 51 C) ; DB933: 5 CTA CTT CAC TGC ATA AAC CAC CAC AAG TAG 3 ( tm 52.4 C) ) ( Fig. 3 2) followed by reverse transcription with reverse primer DB933 and MMLV reverse transcriptase as described above. The reverse transcription was followed by PCR with forward primer DB932 as described above. Following PCR, the dsDNA was separated by agarose gel ( 2 % ) electrophoresis in Tris acetateEDTA (TAE) buffer (pH 7.8) containing ethidium bromide at 100 volts for 60 min. A DNA band ( 447 bp ) amplified by DB932 and DB933 was visualized under UV light The band was cut from the gel eluted with QIAquick gel extraction kit (Qiagen, Germantown, MD) and cloned into pCR2.1 according to manufacturer instructions (Invitrogen, Carlsbad, CA ) INV F E. coli cells were transformed and grown overnight on LB plates in the presence of 100 g/ m L Kanamycin. Colonies were lifted from the plates and grown in LB medium in the presence of Kanamycin and plasmids were purified from each culture with a QIAprep

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38 Spin Miniprep Kit (Qiagen, Germantown, MD). Plasmids were sequenced using a BigDye Terminator v3.1Cycle Sequencing Kit (Applied Biosystems, Foster City, CA) and sequ e nces analyzed by th e DNA sequencing core at the University of Florida ( http://langsat.biotech.ufl.edu/ ) showing that the 447 bp dsDNA obtained by RT PCR from Ae. aegypti RNA is similar to the EST sequence released by TIGR A primer complementary to the poly A tail was synthesized (dT17 adapter DB265 ) (5 GAC TCG AGT CGA CAT CGA TTT TTT TTT TTT TTT TTT TT 3) (tm 67 C ) ( Fig. 3 2) and used as a reverse primer in RT PCR. The sequence between DB932 and DB933 allowed the design and synthesis of a different forward prim er (DB940: 5 TAC ATT GCT GTT GTG CGC AGG ATG 3) ( tm 60.4 C ) ( Fig. 3 2) that is located 350 bp downstream of DB932. After RT PCR using dT17 and DB932 a band corresponding to 681 bp was detected by agarose gel ( 2 % ) electrophoresis in TAEbuffer under a UV light The band was cut and ligated in to pCR2.1 INV F E. coli cells were transformed and the plasmid purified and sequenced as described before. Sequence analysis showed that the 3 end of the jmtA gene all the way to the polyA end was obtained using primers DB265 and DB932 ( Fig. 3 2). To find the 5 end of jmtA Superscript II reverse transcriptase was used (H ttemann 2002). First a mixture of 5 g total RNA, 0.5 L RNase inhibitor (20 U), 5 L dNTPs (2 mM each), 1 L 15 M genespe cific forward primer (DB954: 5 GGA TAG CTG ATC ATA AAT GTC GAA 3) ( tm 52.1 C ) ( Fig. 3 2) was heated to 80 C for 30 min and immediately placed on ice for 2 min. Then 5 L of 5 x first strand buffer (Invitrogen, Carlsbad, CA ), 2.5 L DTT and 1.5 L reverse transcriptase (300 U) were added to the reaction and incubated for 90 min at 37 C The mixture was then heated

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39 for 3 min at 85 C and immediately put on ice for 2 min. Reverse transcriptase ( 1 L; 200 U) was added followed by a 60 min incubation at 37 C The reverse transcriptase was then heat i nactivated for 10 min at 75 C and RNA was degraded by adding 1 L RNase H (2 U) and the reaction incubated for 20 min at 37 C The RNase H was heat inactivated for 10 min at 75 C and the cDNA was purifie d using a QIAquick PCR purification Kit (Qiagen, Germantown, MD). A poly A tailing reaction was then performed by combining 5 L cDNA, 3 L dATP, 4 L 5 x reaction buffer ( Invitrogen), 1 L (14 U) terminal deoxynucleotidyl transferase and sterile distilled water up to a total volume of 20 L The mixture was incubated for 3 min at 37 C and heat inactivated for 10 min at 75 C. Second strand synthesis was done by combining 5 L of the templat e with 2 L MgCl2, 2 L dNTP mix (10 mM of each NTP), 2.5 L 10 x PCR buffer, 1 L reverse primer (DB 954), 1 L forward primer (DB265), 0.25 L AmpliTaq DNA polymerase (2.5 U) and 11.25 L sterile distilled water. PCR was carried out as follows: denaturat ion for 3 min at 95 C (1 cycle), annealing for 4 min at 48 C and extension for 40 min at 60 C (1 cycle each), denaturation at 95 C for 30 seconds annealing for 30 seconds at 48 C and extension for 2 minutes at 60 C (40 cycles ) with a final extension cycle for 15 min at 60 C Since this approach yielded poor DNA sequences at the 5 end a second approach using the Ae. aegypti genome (GenBank accession number AAGE02000000) was used. A gene specific primer at the 5 end was synthesized (D B 972: 5 ATG AAC AAA CCT AAT CTT TAT CAC CGA 3) ( tm 55.7 C ) ( Fig. 3 2) Reverse transcription was performed with Superscript II reverse transcriptase according to the protocol described by H ttemann until the DNA purification step (see above) using a genespecific reverse primer DB954 ( Fig. 3 2). After RT PCR, 5 L of the

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40 reaction mixture was removed and the dsDNA reamplified using a second PCR mixture containing forward and reverse primers DB954 and DB972, respectively ( Fig. 3 2) as follows: denaturation for 3 min at 95 C (1 cycle), annealing for 4 min at 48 C and extension for 40 min at 60 C (1 cycle), denaturation at 95 C for 30 seconds annealing for 30 seconds at 48 C extension for 2 minutes at 60 C (40 cycles), with a final extension cycle for 15 min at 60 C A band of 478 bp was detected under UV after running an agarose gel (2% ) electrophoresis in TAE buffer containing ethidium bromide The band was topocloned into pCR2.1, INV F E. coli cells we re transformed and the plasmid purified and sequenced as described before. Sequencing detected the 5 end of jmtA gene ( Fig. 3 2). A full length cDNA was then amplified u sing forward and reverse primers (DB 972 and DB 933) and overlapping sequences at the 3 end and 5 end of the two partial sequences that were obtained from the mRNA. The full sequence was cloned into pCR2.1, and the plasmid sequenced using cycle sequencing showing a full length sequence of the message from the start to the stop signal ( Fig. 3 3) Expression of jmtA in B acterial C ells JHAMT Expression Ae. aegypti jmtA was cloned into t he expression vector pET Duet 1 (Novagen) ( Fig. 3 4) using forward and reverse primers (DB1015) 5 AAA AAA GGA TCC T ATG AAC AAA CCT AAT CTT TAT CAC CGA 3 ( tm 67.2 C ) and (DB985) 5 AAA AAA GCG GCC GCC T AC TTC ACT GCA TAA ACC AC 3 ( tm 72.7 C ), respectively ( Fig. 3 5). Primers (DB1015) and (DB985) carried a restriction site s for B am HI and N ot I respectively. The amplified jmt A was cloned into pCR2.1 and the cloned gene was cut with B am HI and N ot I purified by agarose electrophoresis and ligated into pET Duet 1 that was opened with the same restriction enzymes and purified by agarose

PAGE 41

41 electrophoresis An extra base (T) was added between the B am HI restriction site and the start codon of the sequence to keep the sequence in frame. jmtA was cloned in the first multiple cloning site of pET Duet 1 downstream of the hexahistidinetag to allow the purification of JHAMT on a Ni column Th e vector was sequenced by cycle sequencing as described above, to confirm the presence of a full length jmt A sequence. T he vector carrying the jmtA was cloned in to E. coli One Shot BL21 (DE3) competent cells, which are genetically engineered to carry a T7 RNA polymerase. Cells were grown overnight at 37 C in LB medium (5 mL) in the presence of ampicillin (100 g/mL). Following incubation, the cells were centrifuged at 4,000 rpm at 4 C for 10 minutes and the pellet resuspended in LB medium (250 mL) containing 200 g/mL ampicillin. The resuspended cells we re then grown at 37 C until an OD600 of 0.4 (~ 4 x 108 cells/mL) was reached. The cells were centrifuged at 4,000 rpm at 4 C for 10 minutes and the pellet resuspended in 500 mL LB medium containing 500 g/mL ampicillin until an OD600 of 0.6 (~ 6 x 108 c ells) was reached. Transcription of jmt A was initiated D 1 thiogalactopyranoside (IPTG) (Fisher Atlanta, USA ), which inhibits the lacI repressor of pET Duet 1 The cells were then incubated overnight at room temperature, shaking at 225 rpm in the presence of IPTG to express JHAMT Extraction of JHAMT from Bacterial C ells After the induction period, the BL21(DE3) cells were incubated with a bacterial extraction reagent BPER (Pierce Rockford, IL) and a bacterial protease inhibit or cocktail (Sigma Aldrich St. Louis, MO ) for 20 minutes at room temperature while shaking. At that time, g lass beads ( 180 m) ( Sigma Aldrich, St. Louis, MO ) were added to the incubation mixture and the cells were broken in a FastPrep Instrument for 20

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42 seconds centrifuged at 14,000 rpm for 10 minutes at 4 C and the cell extract stored in a 5% glycerol solution mercaptoethanol (BME; Sigma Aldrich, St. Louis, MO ) at 20 C Purification of JHAMT The extracted histidine tagged protei n was adsorbed onto a nickel column (NiNTA, Qiagen) that was equilibrated with a wash buffer ( 50 mM NaH2PO4, 300 mM NaCl, pH 8.0). The column was first washed for 100 minutes (20 ml) with the same buffer followed with 15 ml ( 60 min ) washes of imidazole (20 mM, 40 mM and 60 mM) in the wash buffer to elute proteins that nonspecifically bound the column. JHAMT was eluted from the column in 250 mM imidazole after the column was washed for 2 h in 1.0 mL fractions containing 20 mM MgCl2 (final concentration). P rotein concentrations of each fraction were determined at 595 nm by a Bradford protein assay (BioRad CA ). To prevent loss of enzymatic activity during the Ni column purification the wash buffer was fortified with 10% glycerol, 20 mM BME and 1 mM phenyl methanesulphonylfluoride (PMSF; Sigma Aldrich, St. Louis, MO ). Further characterization of the enzyme, however, showed that MgCl2 was not needed for enzymatic activity and in subsequent purifications it was not added to the fractions that were collected during the Ni affinity chromatography. To maintain enzymatic activity, with no loss for several months, glycerol was added to each fraction with JHAMT activity after the Ni affinity chromatography to a final concentration of 50% Fractions after the Ni affinity chromatography, were run on a 10 % SDS PAGE (Laemmli 1970), t he gel was stained for 4 h with Coomassi e Brilliant Blue R250 (Fisher Atlanta, GA ) and destained with methanol/glacial acetic acid (5 % / 7 % ) A band corresponding with JHAMT (Mr 32 kDa) was excised and analyzed by Mass

PAGE 43

43 Spectrometry at the university of Florida Biotechnology protein core (http:/ /www.biotech.ufl.edu/ProteinChem/ ). Molecular Modeling of JHAMT A preliminary threedimensional ribbon model of Ae. aegypti JHAMT was built and the structural energy minimized using SYBYL molecular modeling software. The model was then sent to Drs G. Smag ghe and Pierre Roug at Ghent University and UMR Universit Paul Sabatier respectively, who refined the initial draft model. M ultiple amino acid sequence alignments were carried out with ClustalX (Thompson et al. 1997) using the Rislers structural matri x for homologous amino acid residues (Risler et al. 1 998) The Hydrophobic Cluster Analysis (HCA) ( Gaboriaud et al. 1987) was performed to delineate the conserved secondary structural features (strands of sheet and stretches of helix) along the amino acid sequence of JHAMT by comparison with CbSAMT ( Zubieta et al. 2003) that was used as a model. HCA plots were generated using the HCA server (http://bioserv.rpbs.jussieu.fr). Molecular modeling of JHAMT was carried out on a Silicon Graphics O2 R10000 wo rkstation, using the programs Insight II, Homology and Discover 3 (Accelrys, San Diego CA, USA). The atomic coordinates of the salicylic acid carboxyl methyl transferase (CbSAMT) of the fa i r y fans Clarkia breweri in complex with the substrate salicylic aci d (RCSB Protein Data Bank code 1M6E) ( Zubieta et al. 2003) were used as template to build a three dimensional model for JHAMT In spite of the moderate identity (19.5 % ) and similarity (54.5 % ) that JHAMT shared with the template Cb SAMT, a comparison of their HCA plots indicated a very similar organization of the secondary structural features of both enzymes that allowed us to build an accurate

PAGE 44

44 threedimensional model. Steric conflicts were corrected during the model building proce dure using the rotamer libr ary (Ponder et al. 1987) and the search algorithm of the Homology program (Mas et al. 1992) maintained a proper sidechain orientation. An energy minimization of the final model was carried out by 250 cycles of steepest descent using the cvff force field o f Discover. PROCHECK (Laskowski et al. 1993) was used to assess the geometric quality of the threedimensional model. A bout 67 % of the residues of the model ed JHAMT were correctly assigned to the best allowed regions of the Ramachandran plot (vs 63% for 1 M6E used as a template), the remaining residues are located in other allowed regions of the plot except for eight residues (Gln14, Leu66, Ile99, His111, Lys163, Tyr180, Lys199, Pro226) located in the non allowed region (result s not shown). These misplaced residues are located in loops exposed on the surface of the model. Molecular illustrations were drawn with PyMol (W.L. DeLano, http://pymol.sourceforge.net). The fold recognition program Phyre (http://www.sbg.bio.i c.ac.uk/phyre/html/index.html) (Mayoral et al. 2009) that use s structurally related methyl transfer ase proteins as templates yielded a readily superposable threedimensional model. F ew discrepancies with the shape of the loops connecting the helical stretches and sheets of JHAMT were observed in the model ed structure. T hese discrepancies however, occur outside the ligand binding cavity responsible for the binding of JH III acid Docking of JH III acid to JHAMT was performed with Insight II using Discover 3 as a for ce field. Clipping planes of JHAMT harboring JH III acid in its ligand binding cavity were rendered with PyMol.

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45 Radioactive bioassay for JHAMT Determination of JHAMT Activity HPLC. JHAMT enzymatic activity was measured using a novel radioactive bioassa y. JHAMT activity was measured in a reaction mixture (100 L) containing 50 mM Tris HCl buffer (pH 7.9), 5 % glycerol 3H]S Adenosyl Methionine (SAM) (Perkin Elmer, Waltham, MA) 0.141 M or 600 M SAM (Sigma, St. Louis, MO) and various concentrations of juvenile hormone substrates and JHAMT incubated for different time intervals at 25 C. The reaction was stopped with acetonitrile (ACN) (500 nM MF and 250 nM JH III that serve as carr iers Aliquots (200600 L) were analyzed by HPLC, (Waters Milford, MA) using a C18 reverse phase (RP) column (150 mm x 4.6 mm, 5 HPLC system consists of a binary HPLC 1525 model pump, a UV etector 2487) and an integrator. The chromatographic separation was performed on the column using a linear ACN water gradient (40% to 100 % ) ( Borovsky et al. 1992). Nonradioactively labeled standards were detected at 214 nm. F ractions (1.0 mL) were collect ed and suspended in 4 m L Scintiverse;LC Cocktail (Fisher, Atlanta GA) and analyzed in a liquid scintillation counter (Tracor Analytic, Elk Grove Village, IL) and the radioactivity converted into fmol or pmol of synthesized JH III or MF. JHAMT activity is expressed as fmol/45 min or pmol/45 min and in mol /mol enzyme/min or mmol/mol enzyme/min. Rapid Biphasic Separation ( RBS ). Because HPLC separations and individual fractions analyses to determine JHAMT activi t ies are time consuming processes, a new technique was developed to rapidly analyze multiple reactions Purified JHAMT was incubated with different substrate s in a reaction mixture ( L) containing

PAGE 46

46 SAM (S [methyl -3 5% glycerol and 50 mM Tris HCl, pH 7.5 for 45 minutes at room temperature (25 C) After incubation, methanol (50 L) was added to stop the reaction and the tube was vortexed followed by the addition of NaCl solution (10% ) and hexane (1.0 mL). The mixture was thoroughly vortexed, centrifuged for 1 minute at 14,000 rpm and the hexane upper layer was carefully removed into 4 m L Scintiverse, vortexed and counted in a liquid scintillation counter. All incubations were perfor med in triplicates. A control reaction in which JHAMT was not added was also run. Radioactivity of control samples extracted in hexane was subtracted from reactions that were incubated with JHAMT because SAM (S [methyl -3H]) contains radioactively labeled i mpurities that co extract in the hexane layer with JH III and MF. The results of the biphasic extraction were compared with results that were obtained using RP HPLC separations (see above) to make sure that JHAMT activity is not over or under estimated. Linearity of JHAMT Activity The linear range of JHAMT activity was followed for 5, 10, 20, 30, 45 and 60 minutes in a reaction mixture (100 L) containing 5 % glycerol, 50 mM Tris HCl ( pH 7.5) 12.5 g purified enzyme, 100 M FA and 1.1 Ci [3H] SAM ( 0.141 ) Enzyme activity was followed in triplicates using the RBS (see above). Effect of Metal Ions on JHAMT Activity To check for metal dependency E thylenediaminetetraacetic acid (EDTA), a chelating agent that can sequester metal ions such as Ca2+, Mg2+ and Zn2+, was added at different concentrations to JHAMT and the activity of JHAMT was followed to find out if EDTA reduces JHAMT activity, indicating metal dependency.

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47 To find out if MgCl2, CaCl2 and ZnCl2 stabilize JHAMT as was reported for other proteins (Gurd et al. 1955), Mg2+, Ca2+ and Zn2+ ions (10 mM) were separately incubated with JHAMT and JHAMT activity was followed and compared with JHAMT that was incubated without these ions. Each reaction was repeated 3 times. Substrates specificity of JHAMT JHAMT was incubated with eight potential substrates ( Fig. 3 6) to find out the enzymes specificity. Homofarnesoic acid, JH I (cis/trans/cis), JH I (trans/cis/cis), JH I (cis/trans/trans) and JH I bisepoxide were provided by professor Karl Sl ma (Czech Republic, Prague). JH III bisepoxide and farnesoic acid were provided by Professor G. Prestwich, and JH III was purchased from Sigma (St. Louis, MO). Homofarnesoic acid and farnesoic acid were diluted in dimethylsulfoxide (DMSO) to a concentration of 100 M. JH I acid and JH III acid and JHI and III bisepoxide acid were produced by alkaline hydrolysis in ethanol ( Borovsky & Carlson 1992) and diluted in DMSO to a concentration of 100 M. Briefly, JH methyl ester derivatives (500 g, each) were incubated overnight in a 200 L 0.5 M NaOH ethanol solution at room temperature. The conversion into the acid form was tested by RP HPLC (Borovsky & Carlson 1992 and as discussed above). The incubation continued until most of the JH was converted into its acid derivative. Following incubation, the reaction mixture was purified by C18 RP HPLC, fractions corresponding to the acid derivatives were collected, and ACN evaporated under a fine nitrogen stream. The aqueous mixture was then extracted with hexane (1 mL) in the presence of NaCl. The extract was vortexed, centrifuged at 14,000 rpm at room temperature, the hexane layer removed and the aqueous layer reextracted with hexane as above. The two hexane extracts were combined and evaporated under nitrogen to a final volume of 1 mL. The concentration of the JH acids was determined by

PAGE 48

48 running aliquots on HPLC and comparing peak areas to a HPLC calibration curve that was run with known concentrations of standards. Stock solutions for each acid (100 M) were m ade in DMSO and stored at 20 C until used. Different concentrations of the acids were incubated with 1.65 g purified JHAMT in a 50 mM Tris HCl buffer (pH 7.9) containing glycerol (5% methyl [3H] ), 0.141 for 45 minutes a t room temperature. Each incubation was repeated three times and results are expressed as means S.E.M. The results of these experiments were used to find out the affinity of JHAMT to different substrates. To make sure that the RBS and the HPLC determinations were the same, one reaction from each incubation was also run on C18 RP HPLC. Effect of Incr easing Concentrations of S Adenosyl Lmethionine [methyl -3H] on JHAMT Activity JHAMT activity was measured in the presence of increasing concentrations of SAM ( 0.141, 20, 40, 100, 200, 600 M respectively ) to find out the optimal concentrations of the methyl donor in the reaction mixture. Effect of JH III on JHAMT JHAMT (4.5 g purified enzyme) was incubated with different concentrations of JH III (9.375, 18.75, 37.5, 93.75, 187.5, 375 and 3750 M) in a reaction mixture (100 L) containing 1.1 Ci SAM (S methyl [3H ] ), 0.141 5% glycerol, 50 mM Tris HCl, pH 7. 9 and 500 M FA or 500 M JH III acid. The synthesis of JH III or MF was analyzed by the RBS method. Controls were run without adding JHAMT and were subtracted from reactions containing JHAMT. A third reaction containing SAM (S methyl [3H]) (1.1 Ci; 0.141 M SAM ) 5% glycerol, 50 mM Tris HCl, pH 7.5 buffer was incubated for 45 min with JH III

PAGE 49

49 (1375 M), JHAMT (4.5 g purified enzyme) in the presence of both FA and JH III acid (500 M each). The reaction was stopped with ACN (500 L) containing MF (85.1 nM) and JH III ( 250 nM) and assayed by C18 RP HPLC (as described above) allowing the separation of MF from JH III (Borovsky et al. 1992). Because JH III and MF are equally extracted in hexane, the RBS was not used. Effect of JH III Bisepoxide Acid on JHAMT Activity To fin d out if JH III bisepoxide affects the methylation of JH III acid, JHAMT (4.5 g purified enzyme) was incubated with different concentrations of JH III bisepoxide acid (100, 500, 1000 and 3000 M) in a reaction mixture (100 L) containing 1.1 Ci SAM (S me thyl [3H ] ) 5% glycerol, 50 mM Tris HCl, pH 7.5 and 500 M JH III acid. The reaction was stopped with ACN (500 L) containing MF (85.1 nM) and JH III (250 nM) and the reaction assayed by C18 RP HPLC (as described above) which separat es JH III bisepoxide from JH III (Borovsky et al. 1992). In Vivo Biosynthesis of JH III from JH III acid by JHAMT Synthesis of [3H]JH III Acid [3H]JH III (specific activity 10 20 Ci/mmol) was obtained from Perkin Elmer and converted into [3H]JH III acid (JHA) by incubating [3H]JH III (5 Ci; 0.5 nmol ) with 0.5 M NaOH in ethanol (200 L ) at room temperature (Borovsky & Carlson 1992) ( Fig. 3 7). At 24 h intervals, aliquots (1 L) from the incubation mixture were assayed by HPLC to find the amount of JH III that was converted into JHA. When 80% of the original JH III was converted into [3H] JH III acid the incubation mixture was purified by C18 RP HPLC and fractions containing [3H] JH III acid were collected, combined and dried under a fine nitrogen stream. [3H]JH III acid was resuspended in acetone (100 L) and total radioactivity determined.

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50 In Vivo Biosynthesis of JH III by JHAMT in ligated females Three groups of female Ae.aegypti (20 per group) were collected at different times after adult eclosion and th e blood meal. Females were anesthetized with diethyl ether and ligated with a fine nylon thread between the thorax and abdomen. The abdominal ligation isolated the headthorax complex with the CA from the abdomen containing the ovaries, fat bodies and the gut. [3H] JH III acid dissolved in acetone (0.5 L; 15,000 cpm) was topically applied to the head thorax avoiding any contact with the abdomen of ligated females 1, 2 and 3 days after emergence and 5, 24 and 48 hours after the blood meal. Following applicat ions of [3H]JHA, the ligated female mosquitoes were incubated in a humidified chamber for 3 hours. After the incubation, the headthoraces were cut off from the abdomens and homogenized in 600 L ACN. JH III (150 nM) carrier was added and the homogenate centrifuged for 15 minutes at 12,000 g and at 4 C. The supernatant was analyzed by C18 RP HPLC and assayed for the biosynthesis of [3H]JH III which is correlated, in vivo with JHAMT activity ( Fig. 3 8) (Borovsky et al. 1992, Borovsky et al. 1994) In Vitro Biosynthesis of JH III by JHAMT To determine JH biosynthetic activity of Ae. aegypti 500 females were collected at different times during their gonadotrophic cycle and kept at 20 C. The headthorax complexes and ovaries of females 72 hours PBM w ere separately removed into Eppendorf tubes and incubated on ice containing PBS buffer (pH 7.5), 20 mM BME and 0.1 % protease inhibitor cocktail (Sigma, St.Louis, MO) (1.0 mL) The tissues were then homogenized with a Teflon homogenizer and centrifuged for 5 minutes at 14,000 rpm and at 4 C. The supernatant s were concentrated 4fold to 250 L by centrifugation for 1 h at 4,000 rpm at 4 C in Ultra Centrifugal Filter s device s (Millipore Amicon Biller ica,

PAGE 51

51 MA ) After concentration, glycerol was added to each preparation to a final concentration of 50% to stabilize JHAMT Extracts equivalent to 10 headthoraces or 10 ovaries (10 L ) were incubated in Tris HCl buffer (pH 7.9) with 5 % glycerol, 1.1 Ci [3H ] SAM 0. 1 41 and 500 M JH III acid for 45 minutes at room temperature. After incubation, ACN ( 400 L) containing nonradioactive carriers JH III and MF ( 150 nmol each) was added. Each reaction was centrifuged for 1 minute at 14,000 rpm to minimize clogging of the C18 column and 600 L of each reaction chromatographed by C18 RP HPLC (as described above) and fractions (1.0 mL) collected. Eluates that were collected with the JH III nonradioactive standard were evaporated to 600 L and rechromatographed on C18 RP HPLC to reconfirm the results and to make sure that no radioactively labeled impurities were trapped with the sample and coeluted with the JH III standard during the first HPLC run. RNA Mediated I nterference Studies Synthesis of jmtA dsRNAs T he transcription vector pLITMUS 28i (New England Biolabs) was used t o synthesize dsRNA. DB972 and DB954 ( Fig. 3 2) were used as forward and reverse primers, respectively to obtain the first part of jmtA ( nucleotides 1477 ) DB932 and DB98 5 ( Fig. 3 2) were used as forward and reverse primers, respectively to obtain the last part of jmtA ( nucleotides 364837) PCR reactions (20 L) were carried out as follows: denaturation at 94 C for 3 min (1 cycle), annealing for 30 seconds at 55 C, and extension for 30 seconds at 72 C (40 cycles) with a final extension for 5 min at 72 C (1 cycle) After PCR, dsDNA sequences of 477 bp (first part of jmtA ) and 473 bp (last part of jmtA ) respectively were obtained. These were purified by 2% agaros e gel electrophoresis, the appropriate bands were cut from the gel and purified from the

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52 agarose using QIAquick columns (Qiagen). The purified dsDNA was cloned into pCR2.1 (Invitrogen) following manufacturer instructions ( www.invitrogen.com ) and transformed into E. coli INV F. Transformed E. coli cells were grown in LB medium and plasmids carrying both inserts were harvested using a QIAprep Spin Miniprep Kit (Qiagen). Purified plasmids were cut with Xba I and Hind III (Fisher Atlanta, GA ) and the DNA fragments were purified using 2% agarose gel electrophoresis and QIAquick gel extraction columns (Qiagen). The purified DNA fragments were ligated into an open pLITMUS 28i that was cut with the same restriction enzymes and purified by agarose electrophoresis ( Fig. 3 9) The cloned dsDNAs were amplified by PCR using T7 primer (5 TAA TAC GAC TCA CTA TAG 3) and pLITMUS28i carrying the first and last part of jmtA as template. The dsDNA fragments carrying T7 promoter regions at 5 end of the plus and minus strands were purified by 2% agarose electrophoresis and transcribed by RNA polymerase using HiScribe RNAi Transcription kit (New England Biolabs Ipswich, MA ) The transcribed dsRNA (up to 10 mg/mL) was precipitated in the presence of 5 M ammonium acetate (pH 5.2) and ethanol (100% ), dissolved in DEPC treated water or PBS (pH 7.5) and its concentration determined using DNA quant (Biochrom Ltd., Cambridge, England). Injection of dsRNA into Female Ae. aegypti Three days af ter emergence, sugar fed female mosquitoes were injected using a fine pulled needle between the last two abdominal segments with 0.5 L of (2 g/0.5 L to 7 g/0.5 L) dsRNA (first and last parts of jmtA ) Two days later, the injected females were fed blood on a chicken and 48 hours after the blood meal each female mosquito was dissected and egg development determined.

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53 Feeding Female Mosquitoes dsRNA Injecting mosquitoes is a labor intensive and an invasive procedure. To avoid possible complications that can be caused by injections, female mosquitoes were fed dsRNA. Walsche, Rechav and Timmons independently reported that feeding dsRNA to several species of insects was as effective as injecting them (Walshe et al. 2009 Rechav et al. 1999, Timmons et al. 1998) Using these reports a novel method to feed female mosquitoes dsRNA was developed. Ae.aegypti (20 females) immediately after adult eclosion were removed into a small cage fitted with a capillary tube ( Fig. 3 10) containing 20 L of DEPC treated water, sucrose (3% or 5 % ) and 316 g/L dsRNA (first part of jmtA ). A sucrose solution without dsRNA served as a control. Females were allowed to feed for 4 to 7 days and the capillaries were daily refilled to assure that female mosquitoes fed ad libitum freshly prepared and non degraded dsRNA. After feeding on dsRNA and sucrose, females were removed to a larger cage and were fed blood on a chicken to repletion. At 48 and 72 h after the blood meal, female mos quitoes were dissected under a dissecting microscope and their ovaries examined for egg development. Headthoraces, guts, ovaries and fat bodies were removed into tubes containing Trizol (Invitrogen) for Northern blot analyses. To divert the dsRNA from the crop into the gut, 0.2% BSA was added to the 5% sucrose solution containing the dsRNA. Friend et al. (1989) showed that 17% cellobiose mixed with 3.4% sucrose diverts the sugars preferentially into the midgut (Friend et al. 1989). To develop a method that allowed optimal transport of the dsRNA into the gut, I tested the effect of BSA and cellobiose mixed with sucrose together, or alone.

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54 RNA Mediated Interference Studies on Mosquito Larvae with Long Hairpin (LHP) RNA A long hairpin (LHP) sequence of jmtA ( 333 bp) was synthesized by GenScript (Piscataway, NJ) cloned into pPICZB at the Xho I and XbaI restriction sites and the plasmid sequenced to confirm the incorporation of a full length insert ( Fig. 3 11 and 312). The hairpin sequence contains nucleotides 60 to 180 from jmtA at the 5 end, a small 20 bp fragment for the hairpinloop and a complementary sequence to nucleotides 60 to 180 of jmtA at the 3 end. Pichia pastoris cells were transformed with pPICZB carrying the LHP sequence using Pichia EasyComp Transformation kit (Invitrogen). Following transformation, recombinant cells were selected on Zeocin and cells were grown in Pichia minimal medium (MGY) containing yeast nitrogen base or in Buffered MGY (pH 6.0) (Invitrogen) containing 100 g/mL Zeocin. T he cells were grown to OD600 of 2 6 (1.5 x 108 4.5 x 108 cells/mL) and induced with methanol ( 1 % ) for 24, 48, 72 and 96 h in a minimal medium (MM) or buffered MM (pH 6.0) with100 g/mL Zeocin (Invitrogen). After fermentation, the yeast cells were centrif uged and washed 3 times in sterile water. The cells were diluted in sterile water to OD600 of 20 (1.5 x 109 cells), which is equal to the OD600 of a 2 % solution of empty P. pastoris cells that were cloned with pPICZB without an insert. Sterile tap water (1 00 mL) was added to sterile glass bowls with 20 first instar larvae followed with 4 mL of yeast cells (2% solution). Each experiment was repeated three times and larvae were fed: a. brewers yeast (control) b. P. pastoris cells (control) c. P. pastoris cells transformed with pPicZB without an insert and d. P. pastoris cells expressing LHP RNA. The cells were induced with methanol for 24, 48, 72 and 96 h prior to harvest and addition to the larvae. Larval

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55 growth, development and mortality was monitored d aily until after adult eclosion. A probit analysis was used to calculate 50% pupation time (PT50) in the different groups. Northern Blot s Analys e s of jmtA m RNA RNA was extracted from different tissues during different stages in the life cycle of female Ae. aegypti Extracted RNA (about 30 g) was separated by 1% agarose formaldehyde gel electrophoresis and transferred for 3 h onto a positively charged nylon membrane using NorthernMaxTM Northern Blotting Kit (Ambion, Austin, TX). After transfer, the membrane was baked at 120 C in an oven for 15 minutes. RNA markers (Promega, Madison, WI) were used as standards and the markers were visualized by staining with methylene blue (Maniatis et al. 1982). The membrane, with the transferred RNA was prehybridized for 3 hours at 42 C in a roller bottles oven with Ultrahyb Ultrasensitive Hybridization buffer (Applied Biosystems, Foster City, CA). Four different probes were used to target the mRNAs: a. jmtA b. Ae. aegypti actin (control), c. LHP RNA and d. P. pastoris actin (control) The probes were labeled with [32P] labeled dCTP using RediprimeTM II random prime labeling system (Amersham Pharmacia Biotech Piscataway, NJ) or with DECAprimeTM II kit (Ambion, Austin, TX). Membranes were separately hybridized overnight at 42 C with each probe, and the blots washed 2 times for 10 minutes in 50 mL NorthernMax Low Stringency wash buffer (Ambion) and 3 times for 20 minutes with 60 mL of NorthernMax High Stringency wash buffer (Ambion) at 42 C in a roller bottles oven. Washed membranes were exposed to X ray film at 80 C for 8 to 72 h and the film was developed using Kodak processing chemicals, rinsed in water and dried. Probes were stripped from t he membrane using a boiling solution of 0.1 % SDS solution and membranes scanned with a Geiger counter to confirm that the probes were completely removed.

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56 Figure 31. Multiple sequence alignment (MSA) of JHAMT amino acid sequences of Drosophila melanogaster Bombyx mori and Anopheles gambiae. Similar sequences are in black. Figure 32 Sequencing strategy of Ae. aegypti 1410bp jmtA transcript

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57 Figure 3 3 Schematic representation of jmtA in pCR2. 1. Figure 3 4 Schematic representation of cloning jmtA in pETDuet 1 (Novagen) Figure 35 Cloning strategy of jmtA (start stop) in pET Duet 1 (Novagen)

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58 Figure 3 6 JH acid derivatives used for JHAMT activity Figure 3 7 Chemical conversion of [3H]JH III into [3H]JH III acid by methanolic alkaline hydrolysis

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59 Fig ure 3 8 Synthesis of [3H]JH III by JHAMT from [3H]JH III acid Figure 3 9 Schematic representation of the cloned dsDNAs of jmtA in pLitmus28i

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60 Figure 3 10. Small container with capillary tube anchored by a cotton plug (left ) and female mosquitoes feeding from a capillary tube (right) Sucrose solution mixed with dsRNA (red color) Figure 3 11. Long hairpin (LHP) of jmtA ~ 200 nt ~ 200 nt

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61 Figure 3 12. Schematic representation of LHP cloned in pPicZB showing the cloning restriction sites ( Xho I and Xba I) the AOX1 promoter (P) and Zeocin resistant gene (Zeo)

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62 CHAPTER 4 RESULTS cDNA S equence of jmtA The entire A e aegypti jmtA cDNA sequence was obtained by conducting 5 and 3 rapid amplification of cDNA end (RACE) protocols (Frohman 1993). The fragments were cloned, sequenced and the genomic sequence of putative jmtA identified in GenBank (GenBank AAGE02000000). A full length cDNA sequence was generated and deposited in GenBank (Borovsky et al. 2006, accession number DQ409061) ( Fig. 4 1) and the gene was named jmtA ( JHAMT of Ae.aegypti ) The cDNA sequence is 1410 bp long with 837 bp of an ORF coding for 278 amino acids. Two introns (85 bp and 10,780 bp long) were identified in the genomic DNA sequence including a termination region of 520 bp and a 2000 bp long untranslated region (UTR) that consists of a promoter core with a TATA box at 120 bp and a CAAT box at 163 bp ( Fig. 4 2). The jmtA is directionally opposite to the Ae. aegypti genome, the 5 end of jmtA is the 3 end of the genome ( Fig. 4 2). Expression of jmtA in B acterial C ells Purification of JHAMT pET Duet 1 was opened with restriction enzymes BamHI and NotI and jmtA cloned into the open plasmid at the first multiple cloning site with a 6 x h is tidine tag at the N terminus (see Chapter 3 jmtA expression). The recombinant plasmid with jmtA was cloned into E. coli BL21(DE3) and recombinant cells in 500 mL induced with IPTG (as described above). The recombinant JHAMT was purified on a nickel column washed with low concentrations of imidazole (20, 40 and 60 mM) and the purified protein eluted with 250 mM imidazole (as described above). Fractions (1.0 mL) w ere collected and

PAGE 63

63 their protein concentration was measured using a Bradford protein assay (BioRad, CA) ( Fig. 4 3) SDS PAGE and Mass Spectr a Analyses of JHAMT A e. aegypti JHAMT protein contains 27 8 amino acids with Mr 32 kDa. Recombinant cells expressing JHAMT were broken with BPER (Pierce) and glass beads and the protein purified on a nickel column ( Fig. 4 3 ). Proteins eluted at different imidazole concentrations were separated by SDS PAGE (4 % stacking gel, 10% resolving gel) (Laemmli 1970). After staini ng the gel with coomassie brilliant blue (BioRad Laboratories Richmond, CA) a single protein band at Mr 32 kDa that was eluted from the Ni column with 250 mM imidazole was found ( Fig. 4 4 ). The band was cut from the gel, digested with trypsin and analyzed by mass spectrometry at the University of Florida Biotechnology Center at the Protein Core. Mass spectrometry analysis of the digested protein identified 24 peptides that matched 88% of JHAMT protein sequence ( Fig. 4 5 ), confirming the identity of the protein as JHAMT. Molecular Modeling of JHAMT The model ed JHAMT exhibits a canonical structural organization of SAM dependent methyltransferases ( SAM MTs ). It is made of a globular domain, containing an extended sheet characteristic of all other SAM MTs (Zubieta et al. 2003) and a helical cap domain ( Fig. 4 6 A ). The two domains of JHAMT form a long, narrow active site tunnel that can accommodate JH III acid as a substrate ( Fig. 4 6 B ). JH III acid interact s with JHAMT via the hydrophobic amino acid res idues border ing the JH III acid binding tunnel ( Fig. 4 6 C ). The catalytically active His119 residue, associated with the conserved Trp120 residue, is located at the end of the tunnel and positioned in such a way that the carboxylic group of the docked JH III acid becomes correctly lined up for a

PAGE 64

64 methyl group transfer from a donor ( Fig. 4 6 D ). Another pocket located at the opposite side of the JH III acid binding tunnel, is large enough to accom m odate a SAM molecule as a methyl donor ( Fig. 4 6 B, C, D ). The loop containing the conserved SAM binding region 41 DIGCGSG 47 (Mayoral et al. 2009) occurs at the center of the pocket. All together, predictions from t he structural organization of JHAMT and docking of JH III acid into the JH III acid binding tunnel are in accordance with the measured activity of Ae. aegypti JHAMT. Activity of JHAMT C18 RP HPLC was used to assess the activity of JHAMT The method is based on earlier published work that showed that C18 RP HPLC effectively resolves JH I, II, III, MF JH III acid and JH Bisepoxide in one chromatographic step allowing high recovery and rapid quantization of JH and its metabolites (Borovsky et al. 1992 ). JHAMT methylates the carboxylic moiety of JH III acid by transfer ring a methyl group from the co enz yme, SAM to JH III acid converting it into a methylester Initially it was suggested that in Diptera JHAMT primarily converts FA into MF (Li et al. 2003b Tobe et al. 1985) thus, for the initial characterization of JHAMT FA was used as the preferred subst rate for the enzyme. L) containing [3 ), of JHAMT and a Tris HCl buffer (pH 7.9) for 30 min at room temperature. After incubation, ACN MF ( 85.1 nM ) and JH III ( 250 nM ) was added and an aliquot ( ) chromatographed on a C18 RP HPLC column, the column eluted with a linear gradient of ACN/water (40 % to 100% ) fractions (1 mL) were collected and assayed for radioactivity in a liquid scintillation counter (Tracor Analytic, IL) MF standard was eluted at 33.4 minutes ( Fig. 4 7 ) and radioactively labeled MF eluted at 34

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65 min because it is more hydrophobic than MF ( Borovsky et al. 1992) Incubation o f FA with purified or crude preparation of JHAMT converted it into [3H] MF that was collected at 34 min ( Fig. 4 8 ) Following incubation with JHAMT, FA and [3H ] SAM the reaction mixture was chromatographed on C18 RP HPLC and fractions (1.0 mL) analyzed for radioactivity in liquid scintillation counter. Recovery of MF and JH analogues after the C18 RP HPLC was between 55 to 81 % To be able to compare between the HPLC runs, all the results that were obtained by HP LC were corrected to 100% recovery. The specific activities of the purified and crude extract of JHAMT using FA as a substrate are 490.14 protein and 4.82 w that 101.7fold purification was ac hieved after the Ni affinity chromatography as compared with the activity of the enzyme in the initial crude extract (Table 41) JHAMT Activity Using a Rapid Biphasic Separation (RBS) Running RP HPLC is time consuming and labor intensive, so a faster met hod that uses organic phase separation was developed. After incubation, JH III and MF were extracted by hexane which readily extracts hydrophobic molecules like JH III and MF from aqueous solutions. Radioactively labeled JH III or MF in the hexane layer was counted in a liquid scintillation counter. The validity and accuracy of the hexane extraction method was tested by comparing it with samples that were analyzed by RP HPLC. When JH III acid was incubated for 45 min with purified JHAMT (4 g) in a reaction mixture containing JH III acid (500 M) and 1.1 Ci SAM (S methyl [3H ] ), 0.141 M, and the incubation mixture extracted with hexane, 448.36 34.09 fmol of JH III were synthesized. The same incubation mixture that was incubated for 45 min and analyzed by C18 RP HPLC showed that 538.77 fmol JH III were synthesized.

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66 Incubations with FA in the same reaction mixture and extraction with hexane and analysis by C18 RP HPLC showed that 154.12 6.87 fmol and 140.34 fmol MF were synthesized, respectively ( Fig. 4 9 ). These results indicate that RBS with hexane can be used when FA or JH III acid are used to determine the activity of JHAMT. Linearity of JHAMT A ctivity To find out if the activity of JHAMT was linear with respect to incubation periods, JHAMT was incuba ted with 500 M JH III acid and at different times during the incubation the reaction mixture was extracted with hexane using the RBS and the organic layer was analyzed by a liquid scintillation counter. The results show that JHAMT activity is linear for 4 5 minutes ( Fig. 4 10 A, B ). Therefore, an incubation time of 45 minutes was chosen when purified JHAMT was incubated with FA, JH III acid and several other JH III acid analogues. JHAMT Activity in the Presence of EDTA and Metal Ions JHAMT was incubated wit h 500 M JH III acid and 1.1 Ci [3H]SAM ( 0. 141 M ) or 600 M SAM at pH 7.9 in the presence and absence of EDTA (0 to 50 mM) (as described in materials and methods Chapter 3). After incubation, the reaction mixture was extracted with hexane using RBS (materials and methods) and analyzed for [3H]JH III biosynthesis in a liquid scintillation counter. JHAMT activity was the same in the presence or absence of EDTA, indicati ng that metal ions are not required for JHAMT enzymatic activity ( Fig. 4 1 1 A, B ). Addition of Mg2+, Ca2+ and Zn2+ (10 mM each) to the incubation mixture in the absence of EDTA showed that Mg2+ and Ca2+ did not affect JHAMT activity, on the other hand, Zn2+ inhibited the enzymatic activity 16fold at a concentration of 10 mM. These

PAGE 67

67 results indicate that Zn2+ binds the enzyme causing a conformational chang e that affects its activity ( Fig. 4 1 2 A, B ). Substrate Specificity To find out the substrate specificity of JHAMT the enzyme (1.65 g; 51.5 pmol) was incubated separately with low and saturating concentrations of SAM ( 0.141 M and 600 M, respectively; see Fig. 4 16) and different concentrations of JH III acid, JH I acid (cis/trans/trans), FA Homo farnesoate, JH I acid (cis/trans/cis), JH I bisepoxide acid, JH III bisepoxide acid and JH I acid (trans/cis/cis) ( Fig. 3 6 ) and the methylated products were assayed by the RBS assay (materials and methods) ( Fig. 4 13 A, B ). Each determination was repeated 3 times and reactions with the highest activities were analyzed by HPLC to confirm the results that were obtained using RB S. The results were plotted using the Lineweaver Burk reciprocal plot and Michaelis Menten constants (Km and Vmax) were calculated for each of the substrates tested at low and saturating SAM concentrations of 0.141 M and 600 M, respectively (Tables 42 and 4 3) JH III acid and JH I acid (cis/trans/trans) are excellent substrates (Vmax of 38.90 and 25.37 mol/mol enzyme/min, respectively, for 0.141 M SAM and Vmax of 69.54 and 46.72 mmol/mol enzyme/min for 600 M SAM). Farnesoic acid and JH I acid (cis/tr ans/cis) are good substrates (Vmax of 12.57 and 7.84 mol/mol enzyme/min, respectively, for 0.141 M SAM and Vmax of 13.00 and 14.71 mmol/mol enzyme/min, respectively, for 600 M SAM) Homo farnesoate is a moderate substrate (Vmax of 3.72 mol/mol enzyme/m in and 9.62 mmol/mol enzyme/min, for SAM of 0.141 M and 600 M, respectively) JH I bisepoxide acid, JH III bisepoxide acid and JH I acid (trans/cis/cis) are poor substrates, (Vmax of 1.26, 1.12 and 0.71 mol/mol enzyme/min, respectively, for 0.141 M SAM and Vmax of 0.66, 0.59 and 0.37 mmol/mol enzyme/min respectively, for 600 M SAM)

PAGE 68

68 (Tables 42 and 43). These results indicate that the binding of substrates to the enzyme is specific and depends on substrates chirality. JHAMT can distinguish between the cis/trans/trans, the cis/trans/cis and the trans/cis/cis forms of JH I. JH I acid (cis/trans/trans) is a good substrate whereas JH I acid (trans/cis/cis) is a poor substrate (Tables 42, 4 3 and Fig. 4 13 A, B ). Increase in substrate concentrations above 2000 M caused inhibition of the enzymatic activity with all the tested substrates except the substrates that are poorly methylated by JHAMT at low (0.141 M) and saturating ( 600 M) concentrations of SAM ( Fig. 4 13 A, B ). The inhibition of the enzymatic activity at high substrate concentration is probably due to binding of the substrates to a low affinity bi nding site on the enzyme, causing a conformational change, decrease in the methyl transferase activity and not to low SAM concentrations. Thus, all the Lineweaver Burk reciprocal plots to determine the Michaelis Menten constants (Km) use substrate concentr ations of 0 500 M (Fig. 4 14 A H and 415 A H ). The calculated Michaelis constants (Km) and Vmax for all the substrates at low and saturating SAM concentrations are summarized in Tables 4 2 and 43 The Effect of SAM on JHAMT Activity To find out if the reason for the decrease in JHAMT activity with increasing substrate concentrations is not due to the depletion of the co enzyme SAM, JHAMT was incubated with increasing concentrations of JH III acid (50 to 5000 M) and low and saturating concentrations of SAM (0.141 and 600 M, respectively) ( Fig. 4 16 A, B ). Increasing SAM did not prevent substrate inhibition at high concentrations, though at saturating SAM concentrations the substrate inhibition starts at higher concentrations of JH III acid ( 2000 M JH III acid as compared with 500 M for the lower SAM concentration).

PAGE 69

69 Effect of JH III on JHAMT To find out if JH III has a feedback inhibitory effect on JHAMT activity, increasing concentrations of JH III (0 to 3.7 mM) (Sigma, St. Louis, M O ) were added to a n incubation mixture containing JH III acid or FA (500 M) and JHAMT activity determined using the RBS method (materials and method s). At a concentration of 18.75 M JH III stimulated the methylation of FA but did not affect the methylation o f JH III acid However, at higher concentrations (37.5 M to 3.75 mM) JH III inhibited the conversion of FA into MF by 2.5 to 5fold and the conversion of JH III acid into JH III by 1.25 to 4fold ( Fig. 4 1 7 and 41 8 ). These results indicate that JH III at low concentration may stimulate JHAMT to synthesize MF from FA, whereas the conversion of JH III acid into JH III is down regulated by high concentrations of JH III, indicating that perhaps the preferred biosyntheti c pathway for JH III biosynthesis is from JH III acid and not from MF. When both FA (500 M) and JH III acid (500 M) were incubated in the presence of 1.375 m M JH III, the methylation of both substrates was inhibited by 1.89 and 1.99fold, respectively after analysis by RP HPLC ( Fig. 4 19). Effect of JH III Bisepoxide Acid on JHAMT JH III bisepoxide acid has an 89.5fold lower Km value than JH III acid and thus it binds stronger to JHAMT than JH III acid. However, the Vmax of JH III bisepoxide is 118f old lower than that of JH III acid. To find out if the poor methylation of JH III bisepoxide is because it does not bind properly in the active pocket of JHAMT and thus, cannot be properly methylated ( Fig. 4 6 A D ), JH III bisepoxide acid (0, 100, 500, 1000, 3000 M) was incubated with 500 M JH III acid in the presence of [3H]SAM and JHAMT. Increasing the concentrations of JH III bisepoxide acid, indeed, caused a 10fold

PAGE 70

70 decrease in the synthesis of [3H]JH III ( Fig. 4 20) indicating that JH III bisepoxide acid and JH III acid compete for the same active pocket of JHAMT. Because JH III acid has a lower affinity to the enzyme than JH III bisepoxide acid, methylation of JH III acid decreased by 10fold whereas the methylation of JH III bisepoxide acid to JH II I bisepoxide was very low as shown before (Tables 4 2 and 43 ). In Vivo Biosynthesis of JH III Synthesis of [3H]JH III Acid [3H]JH III (Perkin Elmer, Waltham, MA) was converted into [3H]JH III acid using alkaline methanolic hydrolysis (materials and methods) To check for the rate of synthesis of [3H]JH III acid, aliquots ( 1 L) were removed during the hydrolysis, chromatographed by C18 RP HPLC and radioactivity analyzed in a liquid scintillation counter (Tracor Analytic, Elk Grove Village, IL) Twent y four hours after the incubation, 31.5% of the [3H] JH III was converted into [3H]JH III acid and at 72 h 75.3% of the [3H] JH III was converted into [3H] JH III acid ( Fig. 4 21). At that time, the incubation was stopped, the mixture chromatographed by RP H PLC and fractions containing [3H] JH III acid collected, evaporated and dissolved in acetone ( 100 L) and the concentration of [3H]JH III acid determined. In Vivo Biosynthesis of [3H]JH III from [3H]JH III A cid To follow the activity of JHAMT during the life cycle of female Ae. aegypti 15 groups of female mosquitoes (20 females/group; 3 groups per time point) 1 and 2 days after emergence, 5, 24 and 48 h after the blood meal were ligated with a fine nylon thread between the thorax and the abdomen. [3H]JH III acid in acetone (0. 5 L; 15,000 cpm) was topically applied to the headthorax complex of each mosquito. After a 3 h incubation period, the headthoraces were cut and removed into Eppendorf tubes,

PAGE 71

71 homogenized in ACN, centrifuged and the supernatants chromatographed by C18 RP HPLC, fractions collected and analyzed in a liquid scintillation counter. The conversion of [3H]JH III acid into [3H]JH III is similar after 1 and 2 days of adult emergence (6 7 fmol/20 hea d thoraces ), whereas at 5 h after the blood meal the synthesis dropped to about 5 fmol/20 head thoraces and continued to drop to about 4 fmol at 24 h. The synthesis increased at 48 h to about 5 fmol/20 head thoraces. Although the drop in the conversion of JH III acid into JHIII by JHAMT was 1.75fold lower at 24 h as compared to 2 days after emergence it was statistically significant ( Fig. 4 22). T he fluctuation in the synthesis of JH III by head thoraces may indicate that the enzyme may be down regulated in the CA after the blood meal in female Ae. aegypti ( Fig. 4 2 2 ). In Vitro Biosynthesis of JH III from JH III Acid To find the activity of JHAMT, in vitro 500 headthoraces of adult female Ae. aegypti mosquitoes were removed at different times during the gonadotrophic cycle. The tissues were homogenized, centrifuged and supernatants equivalent to 10 tissues were incubated for 45 minutes in an incubation buffer (50 mM Tris HCl, pH 7.9 and 5% glycerol) containing 1.1 Ci SAM ( S methyl [3H]; 0. 141 M ) and 50 0 M JH III acid and JHAMT activity was determined. Higher JHAMT activity was found in extracts from sugar fed females (46.35 fmol JH III/45 min). The activity dropped 2fold 5 h after the blood meal (20.43 fmol JH III/45 min) and increased 2fold at 24 and 48 h (40.15 and 39.7 fmol JH III/45 min, respectively) and dropped to a minimum at 72 h (15.46 fmol JH III/45 min) ( Fig. 4 2 3 ) These results indicate that although enzymatic activity of JHAMT is found in the head th oraces preparation, the CA, througho ut the gonadotropic cycle, the enzyme is down regulated at 5 h after the blood meal, and up regulated between 24 to 48 h and down regulated at 72 h.

PAGE 72

72 To find out if JHAMT is also synthesized by the mosquito ovary, 500 ovaries were removed 72 h after the blood meal, homogenized and aliquots from the supernatant incubated with [3H] SAM and JH III acid as described above. The ovaries synthesized 14.24 fmol JH III/45 min indicating that mosquito ovaries have an active JHAMT and confirming an earlier report that mosquito ovary can synthesize JH III after the blood meal (Borovsky et al. 1994). RNA Mediated I nterference Studies Injecting dsRNA Three days after adult emergence, sugar fed female mosquitoes were injected with different concentrations of dsRNA in DEPC treated water (first part and last part of jmtA ) Two days after the injections, female mosquitoes were fed a blood meal and 48 hours later ovaries were removed under a dissecting microscope and egg development determined. Because the survival rate of fem ales that were injected with dsRNA was about 47% 400 females were injected and healthy females were selected two days later for blood feeding. The high mortality could be due to unidentified effects of the dsRNA on the mosquitoes, because 8090 % of female mosquitoes that were injected with 0.5 L DEPC treated water survived. Egg development in females that were injected with low a dose of dsRNA (1.5 g) was not inhibited, whereas 2.1, 3.1 and 5 g of dsRNA caused 27 % 45 % and 31% inhibition of egg develop ment, respectively (Table 44 ). Injecting 3.1 g of jmtA dsRNA was most effective in inhibiting egg development. Light microscopy of ovaries removed from mosquitoes that were injected with 3 g dsRNA or with water shows that in some mosquitoes the dsRNA did not have an effect whereas in others it completely inhibited yolk deposition in the oocytes ( Fig.

PAGE 73

73 4 24 A, B ). Because the effect of the dsRNA was variable; in some mosquitoes the oocytes were completely empty and in other partial yolk deposition was observed, the average yolk length of all the treated mosquitoes was calculated and compared with the average yolk length of females that were injected with water ( Fig. 4 25). The results show that the average yolk length of controls that were not injected w ith jmtA dsRNA was 2.5fold significantly longer ( 448.94 m 11.0) as compared to the yolk length of dsRNA injected females (234.72 m 21.9) ( Fig. 4 25). Feeding dsRNA To eliminate mosquito mortality, different concentrations of dsRNA mixed with sucrose BSA and cellobiose were fed by capillary to female Ae. aegypti The feeding continued for several days to maximize the effect of the dsRNA. Feeding mosquitoes 5 % sucrose solution and dsRNA was most effective (Table 45 ). When mosquitoes were fed a sucros e solution containing 6 0 g/ 10 L dsRNA, egg development was inhibited more as compared with the other experimental groups (Table 45 ). Feeding mosquitoes for 7 days sucrose solution containing 6 0 g/ 10 L dsRNA inhibited egg development 72 h after the blood meal in 47% of the females as compared to 25% of egg development after feeding for 4 days (Table 45 and Fig. 4 26). To find out if egg development in the second gonadotrophic cycle can be inhibited with jmtA dsRNA, newly emerged mosquitoes were fed 60 g/10 L dsRNA in a 3% sucrose solution for 4 days. After feeding the dsRNA the mosquitoes were blood fed on a chicken and at 96 hours after the blood meal the mosquitoes were provided an oviposition site. Two days after the mosquitoes laid their egg s they were fed a second blood meal and 48 h later the ovaries were removed and egg development determined. Throughout the experiment the mosquitoes were provided with 60 g/10 L dsRNA in a

PAGE 74

74 3 % sugar solution. Yolk length in females that were fed jmtA dsR NA and two blood meals (285.71 m 22.89) was 1.6fold significantly smaller in the second gonodotrophic cycle than in controls (450 m 13.44) ( Fig. 4 27 and 428 A D ). LHP RNA Mediated Interference of Larvae Long hairpin (LHP) RNA was produced in P pastoris cells by cloning a jmtA LHP dsDNA into pPicZB and transforming P. pastoris KM71H cells. Following transformation, the cells were grown and induced with methanol for 24, 48, 72 and 96 hours to find out the optimal time for LHP RNA production. Dur ing the first experiment no buffered media or antibiotics were used during the fermentation and methanol induction period. First instar larvae that were fed brewers yeast cells (2% solution) (controls ) started pupation on day 8 and the pupation time (PT50) of 50% of the larvae, determine d by probit analysis was at day 10 ( Fig. 4 29 A T able 4 6). First instar larva e that were fed P. pastoris cells transformed with jmtA LHP RNA and induced with methanol for 24 hours, started pupation on day 8 with a PT50 a t day 9 ( Fig. 4 29 B Table 4 6). Larvae that were fed P. pastoris cells transformed with jmtA LHP RNA and fermented for 48 hours started pupation on day 9 with a PT50 at day 13 ( Fig. 4 29 C Table 46). Larvae that were fed P. pastoris cells transformed w ith jmtA LHP RNA and induced with methanol for 72 hours started pupation on day 10 with a PT50 at day 18 ( Fig. 4 29 D Table 46). Larvae that were fed P. pastoris cells transformed with jmtA LHP RNA and induced with methanol for 96 hours started pupation on day 9 with a PT50 at day 21.4 ( Fig. 4 29 E Table 46). These results show that jmtA LHP RNA significantly delays the pupation period for larvae that are fed jmtA LHP RNA. P. pastoris cells expressing jmtA LHP RNA were grown in buffered media containing 100 g/ L Zeocin (Invitrogen) to prevent the cells from excising the cloned

PAGE 75

75 jmtA LHP RNA, and induced with methanol. Four groups of first instar Ae.aegypti larvae were fed brewers yeast (2% solution) as controls. Similarly four g roups were fed P. pastoris cells transformed with jmtA LHP RNA. The cells were induced for 96 hours with methanol (1% ). A significant number of the larvae died (43% ) after they were fed P. pastoris cells expressing jmtA LHP RNA ( Fig. 4 30) whereas few larv ae died in the control group (4% ) that were fed non transformed yeast cells. The majority of the larvae (90 % ) that survived the feeding of P. pastoris cells producing jmtA LHP RNA pupated at day 17 after larval emergence whereas the majority of the larvae (90%) in the control group pupated 8 days after larval emergence. This indicates that jmtA LHP RNA not only delays pupation but is also lethal to the developing larvae. These observations prompted me to test the jmtA LHP RNA transformed P. pastoris cells after different times of induction (24, 48, 72 and 92 hours) with methanol in buffered media (pH 6.0) and in the presence of 100 g / l Zeocin (Invitrogen) for larval survival. Cells that were transformed with an empty parental vector (PV) were induce d for 72 hours as an additional control. Cells that were induced for 72 hours with methanol killed 53 % of the larvae and the majority (90 % ) of the surviving larvae (47 % ) had pupated on day 12 after larval emergence. Cells that were induced for 24 hours caused 30% mortality and the larvae that survived pupated within 10 days of larval emergence, whereas recombinant cells that were induced with methanol for 96 h caused 20% larval mortality and the surviving larvae pupated within 10 days of larval emergence Recombinant cells that were induced with methanol for 48 hours caused 13% mortality and the larvae pupated within 10 days of larval emergence. On the other hand, feeding cells containing PV and Brewers yeast caused low mortalities of 8 % and 5%

PAGE 76

76 respect ively and the surviving larvae pupated within 11 days and 8 days, respectively ( Fig. 4 31). Pupae that were fed on the recombinant cells were kept, and adults that emerged were transferred into cages in order to find out if their egg development is going t o be effected, however, one week after adult emergence 85 % of the adults died. Northern Blot Analys e s Northern blot analyses of jmtA transcripts were done on head thoraces with the brain and the CA, whole abdomens containing the gut and ovaries, on ovari es during the different stages of the gonadotrophic cycle of female Ae. aegypti and on ovaries of females fed with 60 g/10 l dsRNA RNA (30 g) was extracted during the gonadotrophic cycle, separated by agarose formaldehyde gel electrophoresis (1% ) and transferred onto a positively charged nylon membrane and hybridized with jmtA specific probe. High amounts of jmtA transcript were found in the head thoraces (CA) 1 and 2 days after adult eclosion, and 5 hours after the blood meal. The amount of the transcript decreased, reaching a low at 48 and 72 hours after the blood meal where faint jmtA transcript bands were observed ( Fig. 4 32). On the other hand, 1 and 2 days after adult eclosion no jmtA transcripts were found in the abdomen. The amount of the jmt A transcript increased in the abdomens at 48 hours and at 72 hours the transcript became as abundant as in the head thoraces 1 and 2 days after adult eclosion ( Fig. 4 32). To find out if the ovary is the source of the jmtA transcript, Northern blot analyse s were done on ovaries that were removed from sugar fed females 3 days after emergence and 48 and 72 hours after the blood meal. At 3 days after emergence a faint jmtA transcript band was observed, the amount of the transcript increased at 48 h and reached a maximum at 72 h similar to the results that were obtained with abdomens ( Fig. 4 3 3 and 4 3 2 respectively). These results show that the ovary synthesizes jmtA mRNA

PAGE 77

77 confirming the earlier results reported above that the ovary and the CA synthesize biologically active JHAMT 72 h after the blood meal ( Fig. 4 23). To find out if feeding female Ae. aegypti dsRNA degrades jmtA transcript, ovaries from females that were fed for 7 days with 60 g/10 l dsRNA were analyzed by Northern blot analyses. A degraded RNA band was observed which confirms that feeding dsRNA breaks down jmtA mRNA causing 47 % inhibition in egg development ( Fig. 4 34 and Table 45 ).

PAGE 78

78 Figure 41. Nucleotide sequence of jmtA cDNA (Borovsky et al. 2006) ( GenBank accession number DQ 409061). The poly adenylation signal is underlined in black, the SAM binding region is underlined in red. The catalytic amino acids His119 and Trp120 are under lined with a broken line.

PAGE 79

79 Figure 42 G enomic DNA sequence of jmtA including introns (black), termination region and TATA and CAAT boxes on the promoter region. The direction of the gene 5 to 3 (arrow) is opposite to Ae. aegypti genome. Fig ure 4 3 Nickel column purification of JHAMT Protein concentrations were measured in 1 ml fractions using a Bradford protein assay and JHAMT was eluted with 250 mM imidazole (Im.)

PAGE 80

80 Figure 44 SDS polyacrylamide gel electrophoresis of fractions obtained by nickel column purification. WB1 = wash buffer without imidazole, WB2 = wash buffer with 20 mM imidazole, WB 3 = wash buffer with 40 mM imidazole. Elution buffer contained 250 mM imidazole. Figure 4 5 Mass spectrometry of the 32 kDa band excised from the SDS gel. Twenty four unique peptides matching 88 % of the entire protein sequence of JHAMT were found (yellow color) 32 kDa

PAGE 81

81 Figure 46 Molecular modeling of Ae. aegypti JHAMT. A) Ribbon diagram of the modeled JHAMT showing the overall organization of the enzyme. N and C correspond to the N and C terminal ends of the polypeptide chain. B) Inside view of JHAMT showing the central position occupied by His119 (blue stick) and Trp120 (yellow stick) residues at the junction of the JH III acid binding tunnel (red dotted line and arrow) and the methyl donor binding pocket (blue dotted line and arrow). C) Ribbon diagram showing the hydrophobic residues (orange colored sticks) border ing the JH III acid b inding tunnel (indicated by dotted lines). His119 and Trp120 residues are colored blue and yellow, respectively. Cavities accommodating the juvenile hormone acid and the methyl donor are indicated by pink and blue arrows, respectively. D) D ocking of JH III acid (pink colored stick) to JH III acid binding tunnel ( shown by a red dotted line). The JH III acid and methyl donor binding cavities are indicated by a pink and blue arrow, respectively. His119 and Trp120 residues are colored blue and yellow, respectively A

PAGE 82

82 Figure 46. Continued. C B C D

PAGE 83

83 Figure 47 C18 RP HPLC of JHIII and MF standards. Absorbance (AU) was followed at 214 nm. Fig ure 4 8 Conversion of FA into MF by purified JHAMT (blue line) and crude extract (red line) [ 3 H] MF FA MF

PAGE 84

84 Table 41. Purification of JHAMT Figure 49 Synthesis of [3H]JH III and [3H]MF from JH III acid and FA respectively with purified JHAMT (4 g) and 1.1 Ci (0. 141 M) [3H]SAM.

PAGE 85

85 Figure 4 10. Linear relationship between JHAMT activity and time. A) 0. 141 M SAM B) 600 M SAM. E ach point is a mean of 3 independent incubations S.E.M A B

PAGE 86

86 Figure 4 11. Activity of JHAMT in the presence and absence of EDTA A) 0. 141 M SAM, B) 600 M SAM. The results are expressed as means S.E.M. of 3 determinations. A B

PAGE 87

87 Figure 4 12. Activity of JHAMT in different salt solutions. A) 0. 141 M SAM, B) 600 M SAM. The results are expressed as means S.E.M of 3 determinations. B A

PAGE 88

88 Figure 413. Substrate specificity of Ae.aegypti JHAMT in the presence of SAM A) 0.141 M SAM, B) 600 M SAM. Farnesoic acid (red), JH III acid (blue dots), JH I acid (cis/trans/trans) (green), Homo farnesoate (black stripes), JH I acid (cis/trans/cis) ( orange), JH III bisepoxide acid (dark blue), JH I bisep oxide acid (pink dots) and JH I acid (trans/cis/cis) (light blue stripes). The results are expressed as means S.E.M. of 3 determinations. A B

PAGE 89

89 Figure 414. Lineweaver Burk reciprocal plots using low SAM concentrations (0.141 M ) with the following JHAMT substrates: A) JH III acid. B) JH I acid (cis/trans/trans). C) Farnesoic acid. D) JH I acid (cis/trans/cis). E) Homo farnesoate. F) JH I bisepoxide acid. G) JH III bisepoxide acid H) JH I acid (trans/cis/cis). A B

PAGE 90

90 Figure 4 14. Continued. C D

PAGE 91

91 Figure 414. Continued. E F

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92 Figure 414. Continued. G H

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93 Figure 415. Lineweaver Burk reciprocal plots using saturating SAM concentrations (600 M) with the following JHAMT substrates: A) JH III acid. B) JH I acid (cis/trans/trans). C) Farnesoic acid. D) JH I acid (cis/trans/cis). E) Homo farnesoate. F) JH I bisepoxide acid. G) JH III bisepoxide acid. H) JH I acid (trans/cis/cis). A B

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94 Figure 41 5 Continued. C D

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95 Figure 41 5 Continued. E F

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96 Figure 41 5 Continued. G H

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97 Table 42. Michaelis constants (Km) and Vmax for JHAMT substrates in the presence of low concentrations of SAM ( 0.141 M) The results are expressed as means of 3 determinations S.E.M. Table 4.3. Michaelis constants (Km) and Vmax for JHAMT substrates in the presence of saturating concentrations of SAM (600 M). The results are expressed as means of 3 determinations S.E.M.

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98 Figure 416. Activity of JHAMT with increasing concentrations of JH III acid using : A) low SAM concentration of 0.141 M SAM and B) a saturating SAM concentration of 600 M A B

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99 Figure 41 7 Synthesis of MF from FA by JHAMT in the presence of increasing concentrations of JH III (0 3.75 mM). Results are expressed as means of 3 determinations S.E.M. Figure 4 18. Synthesis of JH III from JH III acid by JHAMT in the presence of increasing concentrations of JH III (0 3.75 mM). Results are expressed as means of 3 determinations S.E.M.

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100 Figure 4 19. B iosynthesis of JH III and MF from JH III acid and FA in the prese nce of JH III (1375 M) (blue line) and in the absence of JH III (red line). JH III and MF were analyzed by C18 RP HPLC. Inset shows a HPLC run with JH III and MF standards. Figure 420. Synthesis of [3H] JH III by JHAMT in the presence of increasing concentrations of JH III bisepoxide acid (02000 M). Results are expressed as means of 3 determinations S.E.M.

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101 Figure 4 21. Conversion of [3H] JH III into [3H]JH III acid by alkaline hydrolysis Figure 4 22. In vivo conversion of [3H]JH III acid into [3H]JH III by JHAMT. The results are expressed as means of 3 determinations S.E.M. Significantly different from 2 days after eclosion. PE: post emergence, PBM: post blood meal.

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102 Figure 4 23. In vitro biosynthesis of JH III by extracts of Ae. aegypti headthoraces (HT) and ovaries (OV). The results are expressed as means of 3 determinations S.E.M. Significantly different from sugar fed control. Table 44 E gg development in female Ae.aegypti injected with jmtA dsRNA. Different amounts of jmtA dsRNA (1.5 5 g) were injected into sugar fed female Ae. aegypti and 2 days later females were fed a blood meal and analyzed for egg development 48 hours later. Controls that were injected with water developed their eggs normally

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103 Fi g ure 42 4 Light microscopy of yolk development in ovaries of jmtA dsRNA and water injected Ae. aegypti Mosquitoes were injected with jmtA dsRNA (3 g) or water (control) and fed blood 2 days later. Ovaries were removed from females 48 hours after the blood meal and photographed. A) Ovaries from dsRNA injected females, topovary with undeveloped oocytes, bottom ovary with developed oocytes. B) Ovaries with developed oocytes from female injected with water (control). Figure 425. Average yolk length of jmtA dsRNA injected and water injected (control) female Ae. aegypti The results are expressed as means S.E.M. of 90 determinations. *Si gnificantly different using 2 paired student t test ( p 0.05) B A A B

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104 Table 45 The effect of jmtA dsRNA and other supplements on survival and egg development in Ae. aegypti S =Sucrose solution BSA = Bovine Serum Albumine, C = cellobiose. Figure 426. Light microscopy of undeveloped ovaries removed from female Ae. aegypti 72 hours after the blood meal that was fed jmtA dsRNA for 7 days before the blood meal.

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105 Figure 427. Effect of jmtA ds RNA on egg development after a second blood meal The results are expressed as means S.E.M. Significantly different using 2 paired student t test ( p )

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106 Figure 428. Ovaries and oocytes from mosquitoes fed jmtA dsRNA during the second gonodotrophic cycle. A) Pair of ovaries from a mosquito 48 hours after the second blood meal and after feeding the mosquito jmtA dsRNA from emergence. B) Higher magnification and yolk length of the oocytes in A. C) Pair of ovaries f rom a mosquito 48 hours after the second blood meal and after feeding the female sucrose solution (control) from emergence. D) Higher magnification and yolk length of the oocytes in C. A B

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107 Figure 428. Continued. A C D

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108 Figure 429. Pupation periods for larvae that were fed with A) brewers yeast solution (2%). B) P. pastoris cells expressing jmtA LHP RNA and induced for 24 hours. C) P. pastoris cells expressing jmtA LHP RNA and induced for 48 hours. D) P. pastoris cells expressing jmtA LHP RNA and induced for 72 hours. E) larvae fed with P. pastoris expressing jmtA LHP RNA and induced for 96 hours. The results are expressed as means of 3 determinations SEM. A B

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109 Figure 429. Continued. C D E

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110 Table 46. PT50 of Ae. Aegypti larvae that were fed P. pastoris cells expressing jmtA LHP. Recombinant P. pastoris expressing jmtA LHP RNA were fermented for 2496 h and fed to first instar Ae. aegypti larvae. The 50% pupation times (PT50) were determined by pro bit analyses. Controls were fed brewers yeast. Figure 43 0 S urvival of larvae fed P. pastoris cells transformed with jmtA LHP RNA and with non transformed yeast cells. Control: brewers yeast cells (blue line) P. pastoris transformed cells with LHP and induced f or 96 hours (red line) Results are expressed as means of 4 determinations SEM.

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111 Figure 431. Survival of larvae that were fed jmtA LHP RNA yeast cells Control fed with 2 % brewers yeast (dark blue line). Control fed with parental vector (light blue line). Larvae fed P. pastoris cells induced for 24 hours (red line). Larvae fed P. pastoris cells induced for 48 hours (green line). Larvae fed P. pastoris cells induced for 72 hours (purple line). Larvae fed P. pastoris cells fermented for 96 hours (blue/green line). The results are expressed as means of 3 determinations SEM.

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112 Figure 432. Northern blot analyses of head thoraces and abdomens during different stages of the gonadotrophic cycle of female Ae. aegypti HT: Head thoraces, PE: post eclosion, PBM: post blood meal, abd: abdomen. Figure 433. Northern blot analyses of ovaries 3 days post emergence (PE), 48 h post the blood meal (PBM) and 72 h P BM.

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113 Figure 434. Northern blot analyses of ovaries (OV) 72 h PBM from mosquitoes fed with 5 % sugar solution (control) and mosquitoes fed with 60 g/10 l dsRNA. The RNA is degraded when feeding dsRNA (figure on the left).

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114 CHAPTER 5 DISCUSSION Female Ae. aegypti also known as t he yellow fever mosquito, is an anautogenous mosquito that vectors arbovirus es worldwide with economical and public health importance. Mosquito JH plays a major role in many physiological processes, including larval devel opment and vitellogenesis. C haracterization of JHAMT, which is the ultimate enzyme in the JH biosynthetic pathway is important for future biological control of this disease transmitting vector. High homology of JHAMT sequences from A. gambiae, D. melanogaster and B mori and the release of Ae aegypti genome in October 2005 (GenBank accession number AAGE02000000) made it possible to clone and sequence the cDNA of Ae. aegypti jmtA (Borovsky et al. 2006, accession number DQ409061) The cDNA of jmtA is 141 0 bp long with 837 bp of an ORF coding for 278 amino acids. Recombinant jmtA was cloned into pET Duet 1 (Novagen), E. coli BL21 (DE3) cells were transformed and the recombinant JHAMT was expressed and purified by nickel affinity chromatography. The purifi ed enzyme is prone to denaturation and loss of enzymatic activity during freeze and thaw cycles. To overcome this problem the enzyme was stored frozen in small volumes. However, this did not prevent a slow reduction of the enzyme activity with time. The ex pression vector pET Duet 1 carries an ampicillin resistant gene which inhibits transpeptidase and prevents cell wall formation. Wh en ampicillin fails to completely inhibit transpeptidase, the bacterial cells will survive and multiply slowly (Glover et al. 1995) and colonies without the cloned gene will reduce the production of JHAMT. Therefore, the medium was repeatedly refreshed during the 4 hour fermentation and the concentration of the antibiotics was increased. However, this did not increase the produc tion of the recombinant JHAMT appreciably. When the cells

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115 were grown at room temperature overnight in the presence of IPTG the amount of JHAMT increased several fold. The cells were collected and broken with BPER (Pierce), in the presence of protease inhibitor cocktail and purified on a Ni column. All elution buffers used during the purification of JHAMT contained 20 mM BME and 1 mM PMSF and the enzyme was stored in 50% glycerol BME stabilize s enzymes that contain free sulfhydryl groups during their purification by reducing the sulfhydryl groups and preventing opportunistic disulfide bridges (Borovsky et al. 1974). JHAMT has 7 cysteines ( Fig. 4 1 and 46) that need to be reduced to maintain enzymatic activity. PMSF was added to inhibit serine proteases that might degrade the enzyme during the Ni affinity chromatography ( Pierce, technical resource TR0043.1). Purified recombinant JHAMT stored frozen in 50% glycerol and mercaptoethanol (20 mM ) is stable for several months A molecular model of Ae. aegypti JHAMT was built in collaboration with Drs Guy Smagghe at Ghent University, Belgium and Pierre Roug at UMR Universit Paul Sabatier France. The 3 dimensional ribbon drawing of the enzyme show s the canonical structural organization of SAM dependent methyl transferases. Two domains were identified; a globular domain with 6 sheets (Zubieta et al. 2003) and a n helical cap domain consisting of 9 helices. A long narrow tunnel within the domains serves as JHAMT active site and it is composed of hydrophobic amino acid residues that can interact with the substrates and position them in such a way that they can interact with the catalytic amino acids H119 and W120. A pocket located at the opposite site of this tunnel consists of the loop which contains the SAM binding region so SAM can bind there for its methyl t ransfer. Closer examination of the 3 dimensional model detected a

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116 secondary groove below the main catalytic cavity that may allow juvenile hormone acid analogues to bind non specifically caus ing conformational change and steric hindrance to SAM transfer of its methyl group to a JH III acid substrate that binds in the catalytic groove ( Fig. 5 1). JHAMT was characterized with several potential substrates to find out if the 3dimensional modeling and docking predictions can be corroborated by in vitro studies. Incubating JHAMT with JH III acid, JH I acid (cis/trans/cis; trans/cis/cis; cis/trans/trans), FA, homo farnesoate, JH I bisepoxide acid and JH III bisepoxide acid in the presence of saturating concentrations of SAM, showed that the maximal rate of the methyl transfer (Vmax) to JH III acid was 5.3, 4.7 and 1.5fold higher than for farnesoic acid, JH I acid (cis/trans/cis) and JH I acid (cis/trans/trans), respectively (Table 43). The Vmax of homo farnesoate was 7.2fold lower than JH III acid and the Vmax of JH I bisepoxide acid, JH III bisepoxide acid and JH I acid (trans/cis/cis) were 105.4, 117.9 and 187.9fold lower than for JH III acid, respectively (Table 43). These results clearly show that the binding into the narrow catalytic cavity ( Fig. 4 6 ad, and 51) depends on substrate chirality that allows proper alignment in the catalytic cavity and efficient methyl transfer by SAM ( Fig. 4 6 ad, and 51). Indeed, JH I acid (cis/trans/t rans) is 3.2fold more efficiently methylated than JH I acid (cis/trans/cis) and 126.3fold more than JH I acid (trans/cis/cis) (Table 4 3). It is interesting to note that several substrates show a reverse relationship between their Km and Vmax; e.g. JH I acid (trans/cis/cis) has a Km of 0. 08 0.03 M SEM which is 1835fold lower than JH III acid (146.79 15.02 M SEM; Table 43) indicating a much higher affinity to JHAMT, however, the 188fold lower Vmax of this substrate as compared with JH III acid indicates that the alignment of

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117 JH I acid (trans/cis/cis) in the active cavity does not allow it to be efficiently methylated by SAM. Several reports on the characterization of JHAMT from Samia cynthica ricini B. mori and Ae.aegypti were published (Sheng et al. 2008, Shinoda & Itoyama 2003, Mayoral et al. 2008). The Km of Samia cynthica ricini is 55.3 5.9 M SEM whereas the Km for Ae. aegypti is 146.79 15.02 M SEM (Table 4 3) on JH III acid. These authors used a similar radi oactive assay for JHAMT, however, they have neglected to use controls in which [3H]SAM is incubated without the enzyme and the nonspecific radioactivity that are extracted by the hexane used in the RBS are subtracted. Nevertheless, these results indicate t hat the Km for JHAMT from Samia cynthica ricini and Ae.aegypti using JH III acid are similar. Although no Vmax was reported by these authors, from the substrate saturation curve and the Lineweaver Burk double reciprocal plots a Vmax of 3.2 mmol/mol enzym e/min was estimated, which is in the same range as the Vmax we found (69.54 mmol/mol enzyme/min). Characterization of recombinant JHAMTs of Ae. aegypti and B. mori by mass spectrometry reported very high initial enzymatic activities (Vi) for Ae.aegypti and B. mori JHAMT using JH III acid as a substrate (1.033 0.008 and 0.79 0.06 mol/mol/min SEM, respectively) (Mayoral et al. 2008 Shinoda & Itoyama 2003). It is doubtful that these results are correct because kinetic analysis of enzymes by mass spectrometry is not as sensitive or the method of choice as compared with radioactively labeled substrates. Mass spectrometry requires extraction and extensive purification of samples before analysis, and sample recoveries are usually a problem during the many purifications steps that precede the final analysis. Using [3H]SAM and JH III acid the Vmax of Ae.aegypti (this report) and Samia cynthica ricini (Sheng et al. 2008) JHAMT

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118 was determined as 69.54 4.91 m mol/mol enzyme/min and 3.2 mmol/mol enzyme/min, repectively. These values are many fold lower than the very high Vi rates of 1.033 0.008 and 0.79 0.06 mol/mol/min SEM, reported for Ae.aegypti and B. mori respectively using mass spectrometry (Mayoral et al. 2008 Shinoda & Itoyama 2003). Because the Km values for Ae aegypti and Samia cynthica ricini JHAMT using JH III acid as substrate are close (146.79 15.02 M SEM and 55.3 5.9 M SEM, respectively) it is very unlikely that the published reports using mass spectrometry are correct (Shinoda & Itoyama 2003, Mayoral et al. 2008). The Vmax for the methylation of JH I acid (cis/trans/trans) by Ae. aegypti JHAMT is 46.72 11.52 m mol/m ol enzyme/min whereas in B. mori Vi (initial rate) was reported as 1.8 0.32 mol/mol enzyme/min using mass spectrometry (Shinoda & Itoyama 2003). Similarly, the Vi for methyl transfer by Ae. aegypti and B. mori JHAMT to FA is over estimated at 0.260 0. 009 and 0.48 0.07 mol/mol enzyme/min SEM (Mayoral et al. 2008, Shinoda & Itoyama 2003) using mass spectral analysis as compared with Vmax of 13.00 m mol/mol enzyme/min using saturating SAM concentrations. The over estimation of JHAMT methylation rates i ndicates that mass spectrometry for kinetic studies should be used with caution and the results should be confirmed with another technique preferentially a technique that is sensitive like radioactive analysis or fluorescence. Earlier reports suggested th at in Diptera JHAMT mainly converts FA into MF ( Li et al. 2003b Tobe et al. 1985). Mayoral et al. (2008) reported that recombinant JHAMT is 5 fold more efficient in methylating JH III acid than FA. This report using a sensitive radioactive bioassay confirms these observations showing that JHAMT methylates JH

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119 III acid 5.3 times faster than FA (Table 42 ). Thus, JH III acid is the preferred substrate for JHAMT. Low concentrations of JH III stimulate the methylation of FA by JHAMT, but not JH III acid. The conversion of JH III acid and FA into JH III and MF, respectively is inhibited by high concentrations of JH III ( Fig. 4 17 and 418). When JHAMT was incubated together with JH III acid and FA in the presence of high concentration of JH III (1.375 mM) the conversion of JH III acid into JH III and FA into MF was inhibited. The conversion into JH III was much more pronounced indicating that the preferred biosynthetic pathway of JH III is probably from JH III acid and not from MF ( Fig. 5 2). These results indicate that JHAMT probably has additional binding site(s) that JH III binds when it inhibits or stimulates the activity. Such a mechanism may be useful to down regulate the conversion of JH III acid and FA into JH III and MF by feedback inhibition when JH III concentration is high and female mosquitoes need to maintain homeostasis. Increase in concentrat ions of JH acid analogues above 2000 M caused inhibition of all the JH acid analogues except for JH I bisepoxide acid, JH III bisepoxide acid and JH I acid (trans/cis/cis) ( Fig. 4 14). This may be due to the presence of more than one substratebinding sit e on JHAMT. At low substrate concentration the site with high affinity is occupied and normal Michaelis Menten kinetics are observed. When the concentration of the substrate is high the substrate may bind a secondary site on the enzyme causing conformational change, and lowering enzyme activity (Dixon et al. 1979). The 3 dimensional model of JHAMT shows a secondary groove that allows JH III acid analogues to bind causing conformational change and blocking a proper methyl

PAGE 120

120 transfer by SAM to the substrate that is bound at the narrow catalytic cavity ( Fig. 5 1). The Km studies corroborated these observations showing that the affinity of JHAMT to the JH acid analogues that exhibited lower Vmax is much higher than the JH III acid analogues that exhibited a high Vmax. These analogues bind at the catalytic cavity very strongly but are not oriented properly for a successful methyl transfer from SAM ( Fig. 4 6 b d and 51). When increasing concentrations of JH III bisepoxide acid (Km = 1.64 M) were incubated together with 500 M JH III acid (Km = 146.79 M), the conversion of the latter into JH III was significantly reduced. At concentration of 2000 M, JH III bisepoxide inhibited 89 % of JH III acid conversion into JH III ( Fig. 4 20). JH III acid analogues that show low Km and low Vmax could perhaps be developed into a new generation of insecticides that will prevent JH biosynthesis in larvae and thus prevent them from molting and developing into adults. This kind of approach is currently used to fight HIV infections with protease inhibitors that have high affinities and bind to the HIV viral proteases and prevent the virus from infecting human cells (Tomasselli et al. 1990, Akho et al. 2001). JH titers in Ae. aegypti (Borovsky et al. 1992) do not seem to follow the JHAMT activity patter ns found both in in vivo and in vitro experiments using [3H]JH III acid ( Fig. 4 22 and 423). Although JH titer rises after the blood meal (Borovsky et al. 1992), JHAMT activity in vivo and in vitro decreased at 5 hours. The in vitro and in vivo experiment s show that JHAMT is active in the CA (head thorax complexes) throughout the gonadotrophic cycle of female Ae. aegypti JHAMT is up and down regulated throughout the life cycle, but does not seem to be a key regulatory enzyme in the JH III

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121 biosynthetic pat hway as was suggested by Shinoda & Itoyama (2003) and by Sheng et al. (2008) in the Eri silkworm Samia cynthica ricini Borovsky et al. (1994) showed that JH III was not only synthesized by the CA, but also by the mosquito ovary and that JH biosynthesis by the ovaries was three times higher in blood fed mosquitoes than in sugar fed females. Ovaries that were removed from blood fed females 72 h after the blood meal exhibited, in vitro JHAMT activity (14 fmol/45min/10 ovaries) which supports earlier finding by Borovsky et al. (1994). These results suggest that the ovary is probably synthesizing JH III after the blood and that the ovary synth esizes JHAMT. The role of JH in vitellogenesis in mosquitoes is not fully understood. Hagedorn suggested that after adult eclosion, JH makes the ovaries and fat body competent in preparation for the uptake of vitellogenin synthesized by the fat bodies aft er the blood meal (Hagedorn 1974). JH seem s to have a minor role in the biosynthesis of egg yolk proteins and egg development after the blood meal. A second model was proposed by Borovsky in which JH synergi zes with 20hydroxy ecdysone during vitellogenesi s (Borovsky 1981, Borovsky et al. 1985) To find out if JH III plays an important role in vitellogenesis, RNA mediated interference of jmtA was used to block JH III biosynthesis Injecting mosquitoes with dsRNA ( 3 g) blocked egg development in 45% of the mosquitoes that were injected (Table 43 ; Fig. 4 24 and 425 ) Walshe et al. ( 2009), Rechav et al. (1999) and Timmons et al. (1998) independently reported that feeding dsRNA to several insect species was as effective as injecting them. We developed a technique to feed mosquitoes with dsRNA (first part of jmtA ) using a capillary tube which was daily replaced with a newly refilled capillary containing fresh dsRNA to

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122 minimize breakdown of the dsRNA and to minimize mortality due to injury that can be caused by injection. A landings platform for the females was made out of cotton balls so female mosquitoes could land and easily probe and imbibe the fluid in the capillary. Feeding mosquitoes 60 g/10 l dsRNA solutions immediately after adult eclosion for 7 days inhibited egg development at 72 hours after the blood meal in 47% of the mosquitoes that fed the dsRNA and destroyed jmtA tr anscript in the ovary 72 h after the blood meal ( Fig. 4 3 4 ) Feeding mosquitoes dsRNA for 4 days inhibited egg development in 25% of the mosquitoes (Table 44) Feeding mosquitoes 60 g/10 l dsRNA continuously through the first gonadotrophic cycle and for 48 hours after the second blood meal, caused a significant delay in egg development ( Fig. 4 27 and 428 ). Injection and feeding of jmtA dsRNA affected egg development in female Ae. aegypti after the blood meal. However, not all the females that were injected or fed jmtA dsRNA responded indicating that: a. jmtA dsRNA sequences that were used were not efficient and new sequences should be explored, b. the dsRNA did not reach all the tissues t hat synthesize JHAMT, c. short dsRNA sequences should be tried because they may be able to penetrate into cells more efficiently, and d. long hairpin loops should be tried rather than dsRNA. Female mosquitoes that were injected with the dsRNA were 2 days o ld; at that time JH is not needed for egg development anymore (Lea, 1969). However, the effect on egg development is significant and thus, it is very possible that JH or MF are synthesized after the blood meal and have an additional important role in vitel logenesis as was suggested earlier (Borovsky et al. 1985). Feeding Ae. aegypti larvae P. pastoris cells expressing a long hairpin RNA that targets jmtA caused a significant delay in pupation after the yeast cells were induced

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123 with methanol for 96 hours. On the other hand, 90% of the control larvae that were fed brewers yeast (2% ) pupated 12 days after eclosion, whereas only 15 % of the larvae that were fed Pichia cells expressing a long hairpin RNA against jmtA pupated on day 12. It took 30 days for all of the surviving larvae that were fed P. pastoris cells with LHP RNA to pupate ( Fig. 4 31, e). Development of larvae that were fed recombinant Pichia cells producing jmtA LHP RNA and induced with methanol for 24 hours was similar to the control group fed brew ers yeast, indicating that for maximal effect the induction with methanol needs to be longer than 24 hours. To prevent recombinant jmtA LHP RNA loss the media was fortified with 100 g/ml zeocin (Invitrogen) and buffered. Cells that were grown and induced with methanol for 96 hours in the presence of zeocin caused larval mortality of 43% whereas 96% of the larvae in the control group that were fed brewers yeast survived. A delay in pupation of 11 days was again observed in larvae that were fed yeast cells with LHP RNA as compared with controls that pupated in 8 days ( Fig. 4 32). In another experimen t mortality was even higher (53.35% ) when feeding larvae jmtA LHP RNA that was induced with methanol for 72 hours ( Fig. 4 33). In this experiment, pupation was only delayed by 4 days (90% pupated at 12 days after eclosion, compared with 90% of the control group that pupated 8 days after eclosion). In this experiment induction with methanol for 96 hours caused lower mortality (20% ) as compared with induction with methanol for 72 hours ( Fig. 4 33). The majority of the pupae that became adults (70% ) died withi n a week after emergence. The variability between the feeding experiments was probably caused by variations in the shake flask fermentation process and methanol induction. Using a fermenter that controls methanol concentrations oxygen content and pH can re duce the variations between the different

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124 fermentations. Blocking jmtA expression in the larvae causes mortality and delay in pupation. The larvae that ate jmtA LHP RNA were not able to grow and properly molt, as compared with larvae that were fed non reco mbinant yeast. JH is necessary throughout the larval instars for growth and molt cycles. The mortality observed in the emerged adults could possibly be due to a lack of post emergence maturation of the midgut epithelium in mosquitoes which is regulated by JH ( Rossignol et al. 1982) or for the lack of other tissue specific factor(s) that might be regulated by JH during the larval to adult development. Northern blot analyses showed that jmtA is expressed in the head thoraces (CA) at 1 and 2 days after adult eclosion and at 5 and 24 hours after the blood meal. At 48 hours and 72 hours after the blood meal the amount of jmtA transcript is low indicating that low levels of JH are synthesized by the CA. JmtA transcript was not detected 1 and 2 days after adult ec losion in the abdomen (ovary, gut, malpighian tubules and ovaries). At 48 hours after the blood meal a small amount of jmtA was detected, while at 72 h a large jmtA transcript was detected in the abdomen. This transcript is expressed in the mosquito ovaries. These results confirm that JHAMT activity is present in the ovaries and therefore JH III can be synthesized by the ovaries as was reported earlier (Borovsky et al. 1994a). In summary this report shows that jmtA plays an important role in JH III biosynthetic pathway and it appears that the majority JH III is synthesized from JH III acid and not from FA ( Fig. 5 2). The synthesis of MF by JHAMT is slow and it is possible that MF may play an important role in mosquito physiology, however, that role has yet to be discovered.

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125 Figure 51. View of JHAMT molecule showing JH III Acid binding cavity, SAM binding site and a secondary groove that allow JH III acid molecules to bind non specifically. His119 (blue stick) and Trp120 (yellow stick) residues are also shown. Figure 52 Last steps of the JH III biosynthetic pathway. The proposed main pathway of JH III synthesis is shown (red arrows). The conversion through MF is possibly an alternative minor pathway (blue arrows).

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126 LIST OF REFERENCES Agui, N., W.E. Bollenbacher N.A. Granger, & L.I. Gilbert 1980. Corpus allatum is release site for insect prothoracicotropic hormone. Nature. 285: 669 670. Akaho, E., G. Morris, D. Goodsell, D. Wong, & A. Olson. 2001. A study on docking mode of HIV protease and their inhibitors. J. Chem. Software. 7: 103114. Borovsky, D., & E. Van Handel 1979. Does ovarian ecdysone stimulate mosquitoes to synthesize vitellogenin? J. Insect Physiol. 25: 861865. Borovsky, D., E.E. Smith, & W.J. Whelan 1975. Purification and properties of potato 1,4 D glucan: 1,4D glucan 6(1,4 glucano) transferase: Evidence against a dual catalytic function in amylose branching. Eur. J. Biochem. 59: 615 629. Borovsky, D. 1981. In vivo stimulation of vitellogenesis in Aedes aegypti with juvenile hormone, juvenile hormone analogue (ZR 515) and 20hydroxyecdysone. J. Insect Physiol. 27: 371378. Borovsky, D. 1984. Control mechanisms for vitellogenin synthesis in mosquitoes. BioAssays. 1: 264267. Borovsky, D., B.R. Thomas, D.A. Carlson, L .R. Whisenton, & M.S. Fuchs 1985. Juvenile hormone and 20hydroxyecdysone as primary and secondary stimuli of vitellogenesis in Aedes aegypti Arch. Insect Biochem. 2: 7590. Borovsky, D., & D.A. Carlson. 1992. In vitro assay for the biosynthesis and metabolism of Juvenile Hormone by exposed corpora allata of Aedes aegypti (Diptera: Culicidae). J. Med Entomol. 29: 318324. Borovsky, D., D.A. Carlson, & I. Ujvary. 1992. In vivo and in vitro biosynthesis and metabolism of m ethyl farnesoate, juvenile hormone III and juvenile hormone III acid in the mosquito Aedes aegypti J. Med. Entomol. 29: 619629. Borovsky, D., D.A. Carlson I. Ujvry, & G.D. Prestwich 1994a. Biosynthesis of ( 10R ) juvenile hormone III from farnesoic acid by Aedes aegypti ovary. Arch. Insect Biochem. 27: 11 25. Borovsky, D., D.A. Carlson, R.G. Hancock, H. Rembold, & E. Van Handel 1994b. De novo biosynthesis of juvenile hormone III and I by the accessory glands of the male mosquito. Insect Biochem. Molec. 24: 437444. Christophers, S.L. 1960. Aedes aegypti (L.) the yellow fever mosquito. Cambridge University Press, Cambridge, England. Clements, A.N. 2000. The Biology of mosquitoes: development, nutrition and reproduction, vol. 1. CABI publishing, Oxon, England.

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133 BIOGRAPHICAL SKETCH Evelien Van Ekert was born and raised in a small farmers town, Zogge, in Belgium. She attended Pius X Instituut High School in Zele where she graduated in 2001. Evelien attended Ghent University where she received her bachelors in biology and her masters in biotechnology at the Flemish Interuniversity Institute of Biotechnology in the department of molecular and biome dical research under the supervision of professor P. Brouckaert. In 2005, Evelien moved to Florida to work for one year as a visiting scientist in the Florida Medical Entomology Laboratory in Vero Beach in the lab of professor D. Borovsky. After 6 months s he applied for graduate school to pursue a Master of Science degree at the University of Florida, after which she hopes to achieve a Doctor in Philosophy Degree.