Smooth Muscle Titin
in Embryo Implantation and Angiogenesis
vorgelegt von
Diplom-Ingenieurin
Nora Bergmann
aus Berlin
Von der Fakultät III – Prozesswissenschaften
der Technischen Universität Berlin
zur Erlangung des akademischen Grades
Doktorin der Naturwissenschaften
Dr. rer. nat.
genehmigte Dissertation
Promotionsausschuss:
Vorsitzender: Prof. Dr. Roland Lauster
Berichter: Prof. Dipl.-Ing. Dr. Ulf Stahl
Berichter: Prof. Dr. med. Michael Gotthardt
Tag der wissenschaftlichen Aussprache: 21.5.2010
Berlin 2010
D83
Die Dissertation wurde angefertigt am
Max-Delbrück-Centrum für Molekulare Medizin Berlin-Buch
Arbeitsgruppe von Professor Dr. med. Michael Gotthardt
Neuromuskuläre und kardiovaskuläre Zellbiologie
Robert-Rössle-Str. 10
13125 Berlin
Das Staunen ist eine Sehnsucht nach Wissen.
Thomas von Aquin
Acknowledgements
It is my pleasure to acknowledge the people, who supported me to this day.
I thank Professor Dr. med. Michael Gotthardt for the opportunity to work on a fascinating
project. He not only offered excellent working conditions, but also the freedom of research. I am
most grateful for all the inspiring discussions and his trust during the last years.
Prof. Dipl.-Ing. Dr. Ulf Stahl offered me plenty of time and gave me motivation as well as
support. I am deeply thankful for following up and assisting my research.
For the pleasant working atmosphere and the close cooperation I am much obliged to all the
current and former members of the Gotthardt-lab. My research was also supported by their
expert assistance. I like to thank especially Katharina Rost for her helpfulness and Beate
Goldbrich-Hannig for assisting me with the mouse genotyping. I am also grateful to the Hübner-
lab for the DNA sequencing and the Hecker-lab for the mass spectrometry. Furthermore, I have
enjoyed the opportunity to interact with and learn from members of the Willnow-lab.
My immense gratitude belongs to my parents Elfie and Reinhard Bergmann for their love,
patience, guidance, and encouragement. Without them, I would never have enjoyed so many
opportunities. I am deeply indebted to my brother Ulf Bergmann for all his advice and the
scientific ambition that we share. To my friends I am deeply grateful for their effort and
understanding. I am fortunate for all the support I have experienced during my life.
Table of Contents
IX
Table of Contents
List of Figures ......................................................................................................................... XIII
List of Tables ............................................................................................................................. XV
Abbreviations ........................................................................................................................... XVI
Abbreviations of genes and proteins ...................................................................................... XIX
1Abstract .................................................................................................................................. 1
2Zusammenfassung ................................................................................................................. 2
3Introduction ........................................................................................................................... 4
3.1Structure and Function of Muscle ............................................................................. 4
3.1.1Smooth Muscle .......................................................................................................... 4
3.1.2Striated Muscle .......................................................................................................... 5
3.1.3Muscle contraction .................................................................................................... 6
3.2Titin ........................................................................................................................... 8
3.2.1Titin’s structure and localization within the sarcomere ............................................. 8
3.2.2Titin isoforms ............................................................................................................ 9
3.2.3Titin binding partners .............................................................................................. 11
3.2.3.1Z-disc region proteins ...................................................................................... 11
3.2.3.2I-band region proteins ...................................................................................... 11
3.2.3.3A-band region proteins .................................................................................... 12
3.2.3.4M-band region proteins .................................................................................... 12
3.2.4Titin’s kinase ............................................................................................................ 13
3.3Smooth muscle titin ................................................................................................. 14
3.4Embryo spacing and implantation ........................................................................... 15
3.4.1The female reproductive system .............................................................................. 15
3.4.2Embryo implantation ............................................................................................... 16
Table of Contents
X
3.5Angiogenesis ........................................................................................................... 18
3.5.1Sprouting angiogenesis ............................................................................................ 18
3.5.2Uterine angiogenesis during embryo implantation .................................................. 19
3.6Knockout and knockin technology .......................................................................... 20
3.7Aim of the study ...................................................................................................... 21
4Materials and Methods ....................................................................................................... 23
4.1Materials .................................................................................................................. 23
4.1.1Chemicals ................................................................................................................ 23
4.1.2Enzymes .................................................................................................................. 23
4.1.3Bacterial strain ......................................................................................................... 23
4.1.4Vectors and BAC clones .......................................................................................... 24
4.1.5Kits .......................................................................................................................... 24
4.1.6Oligonucleotides ...................................................................................................... 24
4.1.7Antibodies ............................................................................................................... 27
4.2Methods ................................................................................................................... 28
4.2.1Molecular biology methods ..................................................................................... 28
4.2.1.1DNA preparation .............................................................................................. 28
4.2.1.2Determination of nucleic acid concentration ................................................... 29
4.2.1.3Polymerase chain reaction (PCR) .................................................................... 30
4.2.1.4DNA agarose gel electrophoresis ..................................................................... 34
4.2.1.5Real-Time PCR ................................................................................................ 34
4.2.1.6Generation of the targeting vector DsRedKI ................................................... 36
4.2.2Microbiological methods ......................................................................................... 38
4.2.2.1Generation of competent bacteria .................................................................... 38
4.2.2.2Transformation of bacteria ............................................................................... 38
4.2.3Cell biological methods ........................................................................................... 39
4.2.3.1Gene targeting in mouse ES cells .................................................................... 39
4.2.3.2Formation of embryoid bodies ......................................................................... 39
4.2.3.3Preparation and cultivation of primary smooth muscle cells ........................... 39
4.2.3.4Immunostaining of embryoid bodies and smooth muscle cells ....................... 40
4.2.3.5Cell migration assay ......................................................................................... 41
4.2.4Biochemical methods .............................................................................................. 42
Table of Contents
XI
4.2.4.1Protein preparation ........................................................................................... 42
4.2.4.2Protein quantification ....................................................................................... 43
4.2.4.3Protein gel electrophoresis ............................................................................... 44
4.2.4.4Coomassie staining .......................................................................................... 46
4.2.4.5Western blotting ............................................................................................... 46
4.2.4.6Histology .......................................................................................................... 47
4.2.4.7ELISA .............................................................................................................. 48
4.2.4.8Multiplex bead immunoassay .......................................................................... 49
4.2.5Animal procedures................................................................................................... 49
4.2.5.1Maintenance of the mouse colony ................................................................... 49
4.2.5.2Generation of chimeric mice ............................................................................ 50
4.2.5.3Chimera breeding and generation of the TiEx28DsRed strain ........................ 50
4.2.5.4Staging of the estrous cycle ............................................................................. 50
4.2.5.5Timed mating ................................................................................................... 51
4.2.5.6Collection of serum .......................................................................................... 51
4.2.5.7Tissue harvesting .............................................................................................. 51
4.2.5.8Counting of blastocysts .................................................................................... 52
4.2.5.9Evans Blue staining .......................................................................................... 52
4.2.5.10Angiogenesis study .......................................................................................... 52
4.2.6Statistical analysis ................................................................................................... 53
5Results .................................................................................................................................. 54
5.1The DsRedKI mouse model .................................................................................... 54
5.1.1Cloning of the targeting vector DsRedKI ................................................................ 54
5.1.2Generation of the titin-DsRed knockin mouse strain TiEx28DsRed ....................... 56
5.1.3Verification of the titin-DsRed protein and its localization within the sarcomere .. 59
5.2Characterization of the smooth muscle specific knockout of titin’s kinase region . 63
5.2.1Establishing the smooth muscle specific mouse model .......................................... 63
5.2.2Titin’s kinase region is essential for embryo implantation ...................................... 66
5.2.2.1Normal preimplantational embryo development in titin’s kinase region
deficient mice ................................................................................................... 66
5.2.2.2Impaired implantation affects embryo positioning, number of implantation
sites, and uterine maturation ............................................................................ 67
5.2.3Loss of titin’s kinase region influences arachidonic acid metabolism .................... 71
Table of Contents
XII
5.2.4Aberrant angiogenesis in knockout animals lacking titin’s kinase region ............... 72
5.2.5Alteration of tissue remodeling in titin’s kinase region deficient animals .............. 74
5.2.5.1Expression of Murf1 and Fhl2 depends on titin’s kinase region ...................... 74
5.2.5.2The transcription level of Cox2 and Lpar3 is not affected ............................... 75
5.2.5.3Alterations of structural and ubiquitin-proteasome related proteins ................ 76
6Discussion ............................................................................................................................ 78
7Outlook ................................................................................................................................ 93
8Bibliography ........................................................................................................................ 94
9Supplement ........................................................................................................................ 113
List of Figures
XIII
List of Figures
Figure 1: The contractile unit of smooth muscle. ........................................................................... 4
Figure 2: The sarcomere, the contractile unit of striated muscle. ................................................... 5
Figure 3: Molecular basis of muscle contraction. .......................................................................... 7
Figure 4: Titin isoforms and their domain composition. .............................................................. 10
Figure 5: Titin’s binding partners. ................................................................................................ 11
Figure 6: The murine reproductive system and preimplantational embryonic development. ...... 17
Figure 7: Targeting strategy to insert the red fluorescence protein DsRed. ................................. 55
Figure 8: Verification of the targeting vector DsRedKI. .............................................................. 56
Figure 9: ES cell genotyping by PCR. .......................................................................................... 56
Figure 10: Proper localization of titin-DsRed in the sarcomere. .................................................. 57
Figure 11: Genotyping of TiEx28DsRed animals. ....................................................................... 58
Figure 12: Titin-DsRed knockin mice did not have an obvious phenotype. ................................ 59
Figure 13: Detection of titin-DsRed in heart, quadriceps, and soleus. ......................................... 60
Figure 14: The DsRed located close to titin’s N-terminus did not influence the sarcomeric
structure. ..................................................................................................................... 62
Figure 15: Strategy to delete titin’s kinase region in smooth muscle. .......................................... 63
Figure 16: Activity of the SMMHC cre-recombinase in uterine smooth muscle. ........................ 64
Figure 17: Recombination by the SMMHC cre-recombinase in uterine smooth muscle and the
germline. ..................................................................................................................... 65
Figure 18: Normal estrous cycle length and number of blastocysts in knockout females. .......... 66
Figure 19: Impaired embryo implantation in knockout females. ................................................. 67
Figure 20: Reduced number of implantation sites in knockout uteri implies a defect during the
penetration phase. ....................................................................................................... 68
Figure 21: Delayed development of knockout uteri during the penetration phase. ...................... 69
Figure 22: The phenotype observed in the knockout females was due to the deletion of titin’s
kinase region. .............................................................................................................. 70
Figure 23: Deletion of titin’s kinase region affected arachidonic metabolism. ............................ 71
List of Figures
XIV
Figure 24: Altered angiogenesis and PDGF-B serum levels in knockout mice. .......................... 72
Figure 25: Knockout smooth muscle cells did not migrate in response to PDGF-B. ................... 73
Figure 26: Altered expression of Murf1 and Fhl2 in knockout uteri. ........................................... 74
Figure 27: Loss of titin’s kinase region did not alter the expression of Cox2 and Lpar3. ............ 75
Figure 28: Impaired heat shock response and altered cytoskeleton-related proteins in knockout
uteri. ............................................................................................................................ 77
Figure 29: Binding partners of titin’s kinase region. .................................................................... 88
Figure 30: Map of the targeting vector DsRedKI. ...................................................................... 113
Figure 31: Adult smooth muscle knockout animals did not have an obvious phenotype. ......... 114
Figure 32: Increase in the PDGF-B serum levels was not accompanied by changes in selected
cytokines. .................................................................................................................. 114
Figure 33: LIF, an important factor of embryo implantation, was unchanged in pregnant E3.5
knockout uteri. .......................................................................................................... 115
Figure 34: The transcript level of genes encoding selected calcium-related proteins was
unchanged. ................................................................................................................ 115
Figure 35: Dual view of knockout and control 2D-gel. .............................................................. 116
List of Tables
XV
List of Tables
Table 1: Enzymes. ........................................................................................................................ 23
Table 2: Plasmids and BAC clones used for generation of the targeting vector. .......................... 24
Table 3: Kits. ................................................................................................................................ 24
Table 4: Primers for cloning, sequencing, and genotyping .......................................................... 25
Table 5: Primer/probes for Real-Time PCR analysis ................................................................... 26
Table 6: Primary antibodies used for immunofluorescence staining (IF) and Western blotting
(WB). ............................................................................................................................. 27
Table 7: Secondary antibodies used for immunofluorescence staining (IF) and Western blotting
(WB). ............................................................................................................................. 27
Table 8: Primers, template, and expected sizes of the PCR products for cloning. ....................... 30
Table 9: Primers, PCR conditions, and expected product sizes of the genotyping PCRs. ........... 33
Table 10: Knockout mouse models with an implantation phenotype. .......................................... 85
Table 11: Differentially regulated proteins in E3.5 pregnant knockout uteri. ............................ 117
Abbreviations
XVI
Abbreviations
12-HETE 12-hydroxyeicosatetraenoic acid
2D-gel electrophoresis 2 dimensional gel electrophoresis
APS ammonium persulfate
ATP adenosine triphosphate
BAC bacterial artificial chromosome
bp base pair
BSA bovine serum albumin
bw body weight
cDNA complementary DNA
CHAPS 3-((3-cholamidopropyl)dimethylammonio)-1-propanesulfonate
CIP calf intestinal alkaline phosphatase
CON control
Cre causes recombination
C-terminus carboxy-terminus
Da dalton
ddH2O double distilled water
DEPC diethylpyrocarbonate
DMEM Dulbecco’s modified Eagle Medium
DMSO dimethyl sulfoxide
DNA deoxyribonucleic acid
DNase deoxyribonuclease
dNTP deoxynucleoside triphosphate
DTT dithiothreitol
E embryonic day post coitum
ECL enhanced chemiluminescence
E. coli Escherichia coli
EDTA ethylenediamine tetraacetic acid
EGTA ethylene glycol-bis(2-aminoethylether)N,N,N′,N′tetraacetic acid
ELISA enzyme-linked immunosorbent assay
Abbreviations
XVII
ES cells embryonic stem cells
FCS fetal calf serum
FLP flippase recombination enzyme
FN3 fibronectin-type III
FRT flippase recognition target
HET heterozygous
HRP horseradish peroxidase
ICM inner cell mass
Ig immunoglobulin
IgG immunoglobulin G
IF immunofluorescence
IPG immobilized pH gradient
IS implantation site
IVC individually ventilated cage
KI knockin
KO knockout
LB-A plate Luria Bertani-ampicillin plate
LPA lysophosphatic acid
lox-site locus of crossing-over
mRNA messenger RNA
MARP family muscle ankyrin repeat protein family
NP40 nonylphenyl-polyethylenglycol
N-terminus amino-terminus
PAGE polyacrylamide gel electrophoresis
PBS phosphate buffered saline
PCR polymerase chain reaction
pI isoelectric point
PIPES piperazine-N,N'-bis(2-ethanesulphonic acid)
PMSF phenylmethylsulfonyl fluoride
PVDF polyvinylidene difluoride
Abbreviations
XVIII
RNase ribonuclease
rpm rotations per minute
RT-PCR reverse transcription-PCR
S1P sphingosine-1-phosphate
SDS sodium dodecyl sulfate
SEM standard error of the mean
SMCs smooth muscle cells
SMMHC smooth muscle myosin heavy chain
SOC super optimal broth with catabolite repression
TAE Tris-acetate-EDTA
Taq Thermus aquaticus
TE Tris-EDTA
TEMED N, N, N′, N′-tetramethylethylendiamin
TES n-tris-(hydroxymethyl)-methyl-2-aminomethanesulfonic acid
TESCA TES with calcium
Tris tris(hydroxymethyl)-aminomethane
U unit
uw uterus weight
VAGE vertical SDS-agarose gel electrophoresis
WB Western blotting
WT wild-type
X-Gal 5-Bromo-4-chloro-3-indolyl--D-galactopyranoside
Abbreviations of genes and proteins
XIX
Abbreviations of genes and proteins
Abbreviations are given in italics for genes and in bold letters for proteins.
Alox12e E-12/15-LOX epidermal 12/15-lipoxygenase
Alox15 L-12/15-LOX leukocyte 12/15-lipoxygenase
Ankrd2 ANKRD2 ankyrin repeat domain protein 2
Anxa2 ANXA2 annexin A2
Arpc2 ARPC2 actin related protein 2/3 complex
Cacna1c CACN1C L-type calcium channel
Calb1 CALB1 calbindin 1
Calm1 CALM1 calmodulin 1
Capg CAPG capping protein
Car2 CAR2 carbonic anhydrase 2
Carp (Ankrd1) CARP cardiac ankyrin repeat protein
Ckb CKB brain creatine kinase
Camk2a CAMK2A calcium/calmodulin-dependent protein kinase II alpha
Col6A2 COL6A2 collagen type VI alpha 2
Cox2 COX2 cyclooxygenase 2
Cpla2 (Pla2g4a) cPLA2 cytosolic phospholipase A2
Creb1 CREB1 cAMP responsive element binding protein 1
Darp (Ankrd23) DARP diabetes ankyrin repeat protein
Dll4 DLL4 delta-like-4
Dnaja1 DNAJA1 DnaJ (Hsp40) homolog
Etfdh ETFDH electron transferring flavoprotein dehydrogenase
Fga FGA fibrinogen
Fgf2 FGF basic fibroblast growth factor basic
Fhl2 FHL2 four and a half LIM domain protein
Gcsf (Csf3) G-CSF granulocyte colony-stimulating factor
Hbegf HB-EGF heparin binding epidermal-like growth factor
Abbreviations of genes and proteins
XX
Hnrnpab HNRNPAB heterogeneous nuclear ribonucleoprotein A/B
Hspb1 HSPB1 heat shock protein 1
Il1b IL-1β interleukin-1β
Il6 IL-6 interleukin-6
Il15 IL-15 interleukin-15
IP10 (CxCl10) IP-10 interferon-inducible protein 10
lacZ LacZ -galactosidase
Lif LIF leukemia inhibitory factor
Lpar3 LPA3 lysophosphatic acid receptor 3
Mapk13 MAPK 13 mitogen-activated protein kinase 13
Mapk14 MAPK14 mitogen-activated protein kinase 14
Mcp1 (Ccl2) MCP-1 monocyte chemotactic protein 1
MEx1-2 a titin region M-band region encoded by M-band exons 1 and 2
MEx6 a titin region M-band region encoded by M-band exon 6
Mlp (Csrp3) MLP muscle LIM protein
Murf1 (Trim63) MuRF1 muscle-specific RING finger protein 1
Murf2 (Trim55) MuRF2 muscle-specific RING finger protein 2
Murf3 (Trim54) MuRF3 muscle-specific RING finger protein 3
Mylk MLCK myosin light chain kinase
Myom1 MYOM1 myomesin
N2A a titin region N2A region encoded by exon 104
Nbr1 NBR1 neighbor of Brca gene 1
Ndufs3 NDUFS3 NADH dehydrogenase (ubiquinone) Fe-S protein 3
Nrp1 NRP1 neuropilin 1
p62 p62 sequestosome 1
Pcx PCX pyruvate carboxylase
Pdgfb PDGF-B platelet derived growth factor B
Pdgfrb PDGFR platelet derived growth factor receptor
Pdia4 PDIA4 protein disulfide-isomerase A4
Pparg PPAR peroxisome proliferator-activated receptor
Prkaca PKA proteinase kinase A
Prkca PKC proteinase kinase C
Prkcd PKC protein kinase C
Abbreviations of genes and proteins
XXI
Prkce PKC protein kinase C
Prkg1 PKG protein kinase G
Psmc2 PSMC2proteasome 26S subunit ATPase 2
Rack1 (Gnb2l1) RACK1 receptor for activated C kinase 1
S100a1 S100A1 S 100 calcium binding protein A 1
sAnk (Ank1) sANK1 small ankyrin 1
Sfrs3 SFRS3 splicing factor arginine/serine-rich 3
Srf SRF serum response factor
Tagln TAGLN transgelin
Tcap T-cap Titin-cap
Tek Tie2 tyrosine kinase with Ig-like and EGF-like domains 2
Tnf TNF- tumor necrosis factor-alpha
Tpi1 TPI1 triosephosphate isomerase 1
Vegf VEGF vascular endothelial growth factor
Vegfr1 (Flt1) VEGFR1 vascular endothelial growth factor receptor 1
Vegfr2 (Flk1) VEGFR2 vascular endothelial growth factor receptor 2
Vim VIM vimentin
ZEx3 a titin region Z-disc region encoded by Z-disc exon 3
Abstract
1
1 Abstract
The muscle protein titin is with a molecular weight of up to 3700 kDa the largest protein
discovered so far and after actin and myosin the third most abundant protein in vertebrate
striated muscle. Titin integrates into the Z-disc and M-band spanning the half sarcomere and
forms an elastic scaffold along the myofiber. Titin has been implied in signal transduction,
mechanical stability, and maintenance of sarcomere structure in health and disease. While titin in
striated muscle has been well studied, its function in smooth muscle remains unclear although
smooth muscle titin has been described on transcript as well as on protein level.
To study smooth muscle titin, we established a knockin mouse model, in which titin’s N-
terminus was labeled using the fluorescence protein DsRed. Striated muscle analysis of titin-
DsRed mice confirmed the integration of DsRed within titin at the Z/I-junction of the sarcomere.
This modification did not cause any obvious phenotype such as altered viability, heart weight to
body weight ratio, or integration of titin into the sarcomere. The use of the DsRed enables live
cell imaging as well as DsRed detection and makes the knockin mouse a suitable tool to
visualize smooth muscle titin. It is of interest to localize titin in uterus as we were able to
address a functional role for smooth muscle titin in early pregnancy. Titin's M-band exons 1 and
2, which encode titin’s kinase region, were efficiently removed in uterine smooth muscle of a
conditional knockout model using a cre-recombinase. The majority of pregnant knockout
animals showed mislocalization and crowding as well as a reduced number of embryo
implantation sites. Furthermore, knockout mice had a delayed uterine development in early
pregnancy and impaired angiogenesis, which is a hallmark of embryo implantation. The
increased PDGF-B serum levels in pregnant knockout animals and the migration failure of
knockout smooth muscle cells in response to PDGF-B indicate that impaired angiogenesis is due
to a defect in recruiting smooth muscle cells to the site of vessel formation so that newly formed
blood vessels were not stabilized properly. Loss of titin’s kinase region also affected arachidonic
acid metabolism leading to a reduction of its metabolite 12-HETE in pregnant uterus. Our results
demonstrate that signaling of titin’s kinase region via PDGF-B and 12-HETE influences female
fertility most likely by affecting angiogenesis. The increased knowledge of regulatory
mechanisms of angiogenesis during implantation enables novel therapeutic advances for select
forms of human implantation failure and recurrent spontaneous abortion.
Zusammenfassung
2
2 Zusammenfassung
Das Muskelprotein Titin ist mit einem Molekulargewicht von bis zu 3700 kDa das größte
bekannte Polypeptid. Es ist nach Aktin und Myosin das dritthäufigste Protein im quergestreiften
Muskel. Titin integriert in die Z-Scheibe und die M-Bande und überspannt damit das halbe
Sarkomer. Es formt ein elastisches Gerüst entlang der Muskelfaser und ist sowohl an
Signaltransduktion als auch mechanischer Stabilität und Erhaltung der Sarkomerstruktur
beteiligt. Während Titin im quergestreiften Muskel bereits eingehend untersucht wurde, ist seine
Funktion im glatten Muskel unbekannt, obwohl Titin im glatten Muskel auf Transkriptions- und
Proteinebene beschrieben ist.
Um Titin im glatten Muskel zu untersuchen, haben wir ein Knockin-Mausmodell etabliert, in
dem der Amino-Terminus von Titin mit Hilfe des Fluoreszenzprotein DsRed markiert wurde.
Die Analyse von quergestreiftem Muskel der Titin-DsRed-Mäuse bestätigte die DsRed-
Integration in Titin am Übergang von der Z-Scheibe zur I-Bande des Sarkomers. Diese
Modifizierung verursachte keinen offensichtlichen Phänotyp hinsichtlich Lebensfähigkeit,
Herzgewicht und Einbau von Titin in das Sarkomer. Die Verwendung des DsReds ermöglicht
Live-Cell-Imaging und die DsRed-Detektion, so dass unser Knockin-Mausmodell ideal geeignet
ist, um Titin im glatten Muskel zu lokalisieren. Von besonderem Interesse ist Titin im Uterus, da
wir Titin im glatten Muskel erstmalig eine funktionelle Rolle während der Schwangerschaft
zuweisen konnten. Dazu haben wir ein konditionelles Knockout-Modell verwendet. In diesem
wurden Titins M-Banden Exons 1 und 2, die die Titin-Kinase-Region codieren, mit Hilfe einer
Cre-Rekombinase effizient im glatten Muskel des Uterus entfernt. Die Embryo-
Implantationsstellen der meisten schwangeren Knockout-Tiere waren fehllokalisiert, lagen zu
dicht oder waren in der Anzahl reduziert. Zudem war die Entwicklung des Uterus zu Beginn der
Schwangerschaft bei Knockout-Mäusen verzögert. Angiogenese, die bei der Embryo-
Implantation eine wichtige Rolle spielt, war beeinträchtigt und die PDGF-B-Konzentration im
Serum von schwangeren Knockout-Tieren war erhöht. Glatte Muskelzellen aus Knockout-Tieren
zeigten nicht die erwartete Migration nach PDGF-B-Stimulation. Diese Ergebnisse legen nahe,
dass die verminderte Angiogenese durch einen Migrationsdefekt der glatten Muskelzellen zu
dem Ort der Gefäßneubildung bedingt ist, so dass neu gebildete Blutgefäße nicht ausreichend
stabilisiert werden konnten. Durch den Verlust der Titin-Kinase-Region wurde ebenfalls der
Arachidonsäurestoffwechsel beeinträchtigt. Dies führte zu einer verminderten Konzentration des
Metaboliten 12-HETE im Uterus während der Schwangerschaft. Unsere Ergebnisse zeigen, dass
sich die Signaltransduktion der Titin-Kinase-Region via PDGF-B und 12-HETE auf die
Zusammenfassung
3
weibliche Fruchtbarkeit auswirkt, vermutlich durch die Beeinflussung von Angiogenese. Das
Verständnis von regulatorischen Mechanismen der Angiogenese während der Embryo-
Implantation wird neue therapeutische Maßnahmen für einige Arten von Implantationsdefekten
und Fehlgeburten ermöglichen.
Introduction
4
3 Introduction
3.1 Structure and Function of Muscle
Muscles are responsible for contraction that results from thin and thick filaments sliding against
each other. Two types of muscles are classified: smooth and striated muscles. The latter is
subdivided into skeletal and cardiac muscle.
3.1.1 Smooth Muscle
Smooth muscles surround visceral and vascular organs such as the intestine, blood vessels, and
the uterus. The contraction and relaxation of smooth muscles helps to carry food along the
gastrointestinal tract, controls the diameter of blood vessels, and is responsible for embryo
spacing and parturition during pregnancy.
A smooth muscle consists of elongated spindle-shaped cells that have a single nucleus and are
packed with thin and thick filaments (Bagby, 1986). The ratio of thin to thick filaments in
smooth muscle (~20:1) is higher than in skeletal muscle (~6:1; Herrera et al., 2004). The thin
filaments are composed of actin, tropomyosin, caldesmon, as well as calponin and the thick
filaments consist of myosin. Thin filaments are grouped around a thick filament (Figure 1) that
is thereby held in the middle of the smooth muscle contractile unit (Draeger et al., 1990;
Hodgkinson et al., 1995).
Figure 1: The contractile unit of smooth muscle. The schematic representation of three contractile units shows
thin filaments (red) that are grouped around a thick filament (green). The thin filaments are anchored in dense
bodies (yellow) in the cytoplasm and in attachment plaques (brown) at the cell membrane (black).
Introduction
5
The thin filaments are attached to dense bodies in the cytosol or to attachment plaques that are
located at the cell membrane of smooth muscle cells (Geiger et al., 1981; Small and Gimona,
1998). Dense bodies and attachment plaques serve the same function as Z-discs in striated
muscle and consist mainly of the actin-binding protein -actinin. The attachment plaques
additionally contain the protein vinculin that binds to -actinin. In comparison to striated
muscle, in which the contractile units are highly ordered aligned, the contractile units of a
smooth muscle cell are oriented in oblique angles. The lack of visible cross striations leads to the
name smooth.
3.1.2 Striated Muscle
Both skeletal and cardiac muscle cells are striated. Skeletal muscles are attached to the skeleton
by tendons and responsible for body movement. Cardiomyocytes are specialized cells within the
myocardium of the heart and account for the rhythmic contractions to pump blood through the
vessels.
Skeletal and cardiac muscle cells have a similar organization. They are composed of cytoplasmic
myofibrils that run in parallel throughout a multinucleated cell and are attached to the cell
membrane. The myofibrils are made up of sarcomeres (Figure 2), which comprise thick
filaments consisting of myosin and thin filaments consisting of actin, tropomyosin, and troponin.
Figure 2: The sarcomere, the contractile unit of striated muscle. Schematic representation of the sarcomere that
extends between two Z-discs and consists of thin (red) and thick (green) filaments. Titin (black) is integrated with
its N-terminus in the Z-disc and with its C-terminus in the M-band spanning the I-band and A-band. It is linked with
the myosin binding protein C (light green) to the thick filament. At the Z-disc and M-band, opposite titin molecules
overlap so that titin forms a continuous filament along the myofibril. One proposed substrate of the titin kinase
(black boxes) is T-cap (blue circles). Adapted from Gregorio et al., 1999.
Introduction
6
The sarcomere is the contractile unit of striated muscle and defined as the region between two
neighboring Z-discs. Z-discs are the attachment sites for thin filaments of adjacent sarcomeres
that are cross-linked by -actinin. In the middle of the sarcomere is the M-band, in which
myomesin as well as the M-protein cross-link the overlapping thick filament system. The area
nearby the Z-discs, which contains thin filaments, is called the I-band. The thin filaments extend
into the A-band, the region were thick filaments are present. I- and A-band are named for their
properties under polarizing light. The I-band is isotropic leading to a bright appearance whereas
the A-band is anisotropic leading to a dark appearance. The highly ordered structure and the
parallel arrangement of the filaments in the sarcomeres lead to the striated appearance of skeletal
and cardiac muscle.
3.1.3 Muscle contraction
Contraction of smooth as well as striated muscle depends on ATP-dependent cyclic interactions
between thick filaments and thin filaments under influence of cytosolic calcium. This is
described as the sliding-filament model (Hanson and Huxley, 1953; Huxley and Niedergerke,
1954; Huxley, 1957; Guilford and Warshaw, 1998).
In both striated and smooth muscle, ATP is hydrolyzed to ADP by the ATPase activity of the
myosin head. The released energy leads to a conformational change in myosin displacing the
myosin head (Figure 3). Afterwards, the myosin head binds to the actin filament resulting in the
release of ADP and Pi that triggers a power stroke, in which myosin returns to the initial
conformation by the released energy. Thereby, the actin filaments are pulled along the thick
filaments towards the center of the sarcomere and the contractile unit shortens. Binding of ATP
to the myosin head leads to its detachment from actin and the cross-bridge cycle starts again.
Muscle contraction ends if the intracellular calcium concentration is returned to the resting value
by transporting calcium with the help of Ca2+-ATPases and sodium-calcium exchangers back
into the sarcoplasmic reticulum and outside of the cells, respectively. As a result, tropomyosin
blocks again the binding sites of myosin at the actin filaments in striated muscle. In smooth
muscle, the myosin light chain kinase (MLCK) is deactivated and the myosin regulatory light
chain is dephosphorylated by the myosin light chain phosphatase.
Introduction
7
Figure 3: Molecular basis of muscle contraction. If the intracellular calcium concentration is high and the
myosin-binding sites on actin are exposed, ATP is hydrolyzed by the ATPase activity of the myosin head (top left)
leading to a displacement of the myosin head (top right). Binding of the myosin head to actin (bottom right) results
in ADP and Pi release that triggers a power stroke, in which myosin turns to the initial conformation (bottom left).
Thereby, the actin filaments are pulled along the thick filament so that the contractile unit shortens. Binding of ATP
to the myosin head leads to the detachment of the myosin head and the cross-bridge cycle starts again. Modified
from Alberts et al., 2002.
Contraction is controlled by the somatic nervous system in skeletal muscle and by the autonomic
nervous system in heart and smooth muscle. In addition, receptor activation by growth factors
and hormones, which is independent of membrane potential change, also leads to smooth muscle
contraction. The action potential triggers the opening of calcium channels in the muscle cell
membrane and in the sarcoplasmic reticulum so that calcium is released into the cytosol. In
smooth muscle cells, the sarcoplasmic reticulum is only poorly developed so that the increase in
cytosolic calcium in response to an action potential mainly depends on the membrane calcium
channels. As a consequence, change in cytosolic calcium level occur much slower in smooth
muscle than in striated muscle allowing the smooth muscle to produce slow and steady
contractile tension. In striated muscle, binding of calcium to troponin leads to a slight movement
of tropomyosin so that the myosin-binding sites on actin are exposed. This makes binding of the
myosin heads of the thick filament possible. Smooth muscle lacks the tropomyosin-troponin
system that is necessary for rapid response. Instead, intracellular calcium together with
calmodulin binds to the MLCK, which is then able to phosphorylate the myosin regulatory light
chain resulting in myosin head ATPase activity. In contrast, myosin regulatory light chain
phosphorylation only modulates the contractility in striated muscle (Persechini and Stull, 1984).
Introduction
8
3.2 Titin
Titin was first described in 1977 as an elastic muscle protein by the name of connectin
(Maruyama et al., 1977). Two years later, a protein of striated muscle with a megadalton size
was accumulated by electrophoresis (Wang et al., 1979). Because of its large size, the protein
was named titin after the Titans from the Greek mythology. Later it was discovered that titin and
connectin are identical (Maruyama et al., 1981).
3.2.1 Titin’s structure and localization within the sarcomere
Titin is with a molecular weight of up to 3700 kDa the largest protein in mammals and forms
beside actin and myosin the third filament system in striated muscle (Labeit et al., 1992; Labeit
and Kolmerer, 1995). One titin molecule with a length of >1 m spans over a half sarcomere
(Nave et al., 1989) integrating its N-terminus into the Z-disc and its C-terminus into the M-band
(Figure 2). Opposite titin molecules overlap in the Z-disc as well as in the M-band forming a
continuous filament system along the myofibril (Figure 2; Gregorio et al., 1998; Obermann et
al., 1997).
~90% of titin’s protein mass comprises of immunoglobulin (Ig) and fibronectin-type III (FN3)
domains and ~10% are 17 unique domains (Figure 4; Labeit et al., 1990; Labeit and Kolmerer,
1995). ~80 kDa of titin’s N-terminus is integrated into the Z-disc of the sarcomere (Gregorio et
al., 1998). This Z-disc region of titin contains nine Ig domains (Z1-Z9) separated by unique
sequences and up to seven Z-repeats (Labeit and Kolmerer, 1995; Gautel et al., 1996). The I-
band region of titin is 800 to 1500 kDa long depending on the isoform. It consists of tandem
arranged Ig domains and a PEVK region that is rich in prolin (P), glutamate (E), valin (V), and
lysine (K). Furthermore, the I-band region of titin contains the N2A region and the cardiac N2B
region (Labeit and Kolmerer, 1995). The Ig domain regions, the PEVK, and the N2B region are
responsible for titin’s elastic properties. During muscle stretch, first the Ig domain regions are
extended at lower forces by straightening their inter-domain linkers followed by extension of the
PEVK region as well as the N2B region that extends at higher forces (Trombitas et al., 1998;
Helmes et al., 1999; Kellermayer et al., 1997). Since titin forms an elastic connection between
the Z-disc and the M-band, it is responsible for the elasticity of the sarcomere and muscle
passive tension by acting as a molecular spring (Linke et al., 1996, 1999; Granzier et al., 1997;
Trombitas et al., 1999; Watanabe et al., 2002). The A-band of titin accounts for a mass of
~2000 kDa. It has a regular Ig-FN3 super-repeat structure that causes stiffness (Labeit et al.,
Introduction
9
1992; Labeit and Kolmerer, 1995). The ~200 kDa C-terminus of titin is located within the M-
band of the sarcomere and designated as titin’s M-band region (Labeit and Kolmerer, 1995). It
contains a kinase domain and 10 repetitive Ig domains (M1-M10), which are separated by
intervening sequence (is) stretches of varying length (is1-is7; Gautel et al., 1993).
3.2.2 Titin isoforms
The gene locus of titin was mapped for human and mouse to chromosome 2 (Muller-Seitz et al.,
1993; Pelin et al., 1997). The human titin gene consists of 363 exons predicted to code for a
polypeptide with a maximum molecular weight of 4200 kDa (Bang et al., 2001a).
Titin’s A- and M-band domains are constitutively expressed except for M-band exon 5, which is
alternatively spliced (Kolmerer et al., 1996). In contrast, titin’s elastic elements of the I-band
segment are alternatively spliced to determine the elasticity of sarcomeres thereby modulating
passive muscle stiffness (Cazorla et al., 2000; Freiburg et al., 2000). The three major titin
isoforms N2A, N2B, and N2BA (Figure 4) are differently expressed during muscle development
and in different muscles (Labeit and Kolmerer, 1995; Warren et al., 2004; Ottenheijm et al.,
2009b). N2A isoforms are the longest titin molecules with a molecular weight up to 3700 kDa.
One N2A isoform, such as in the soleus, or two N2A isoforms, such as in quadriceps, can exist
in a skeletal muscle type (Ottenheijm et al., 2009b). They contain a large PEVK region and
depending on the muscle type up to 90 tandem Ig domains to increase muscle elasticity (Labeit
and Kolmerer, 1995; Freiburg et al., 2000). Compared to the N2A isoform, the two cardiac titin
N2B and N2BA isoforms are less elastic and shorter with 2970 kDa and 3300 kDa, respectively.
The main titin isoform in the heart is with ~70% N2B (Neagoe et al., 2002). It contains 37
tandem Ig domains and a short PEVK region, which makes the heart the stiffest striated muscle.
The N2B and the N2BA isoforms contain the heart specific N2B domain. The N2BA isoform,
which is predominantly expressed during embryogenesis, contains structural elements of both
the N2B and the N2A isoform varying in the length of the tandem Ig domains and the PEVK
region. Since the N2BA isoform is more elastic than the N2B isoform, the elasticity of the
sarcomeres in the heart can be determined by variation of the N2B:N2BA isoform ratio (Labeit
and Kolmerer, 1995; Cazorla et al., 2000; Freiburg et al., 2000; Bang et al., 2001a).
Introduction
10
Figure 4: Titin isoforms and their domain composition. Titin isoforms spanning over the Z-disc, I-band, A-band,
and M-band of the sarcomere contain Ig domains (red), FN3 domains (white), Z-repeats (green), the PEVK region
(yellow), unique sequences (blue), and the kinase domain (black). The Ig domains Z9 and I1 of titin’s Z-disc to I-
band junction are indicated. A) Splicing of the I-band leads to the N2A isoform with the N2A region in skeletal
muscle and to the shorter N2B isoform with the N2B region in the heart, in which additional the N2BA isoform is
expressed (not shown). B) The novex isoforms 1, 2, and 3 are also obtained by splicing of the I-band in heart and
skeletal muscle, whereas novex 3 contains an alternative titin C-terminus. Adapted from Bang et al., 2001a.
In heart and skeletal muscle additional isoforms are produced by alternative splicing of the Z-
disc and I-band region of titin. Splicing of titin’s central Z-disc region determines the Z-repeat
number (Gautel et al., 1996). Since this Z-repeat is responsible for the titin--actinin cross
bridges, the mechanical properties of the Z-disc are influenced by variation of titin’s Z-repeat
number (Sorimachi et al., 1997). Titin’s Z-disc region is not only expressed differentially in
skeletal and cardiac muscle, but also during embryonic development (Sorimachi et al., 1997).
Titin’s I-band region is also spliced resulting in the novex isoforms 1, 2, and 3 (Figure 4). The
short novex 3 isoform (700 kDa), which contains an alternative titin C-terminus, mediates the
insertion of the long titin isoforms into the sarcomere. The function of the novex isoforms 1 and
2 that contain unique exons located N-terminal to the N2B region is still unknown (Bang et al.,
2001a).
Introduction
11
3.2.3 Titin binding partners
Figure 5: Titin’s binding partners. Proteins that bind to titin’s Z-disc (actinin, obscurin, sANK1, T-cap), I-band
(N2B region: B-crystallin, FHL2, PKA; N2A region: ANKRD2, CARP, calpain 3, DARP, myopalladin; PEVK
region: actin, S100A1), A-band (myosin, myosin binding protein C), and M-band region (calmodulin, calpain 3,
FHL2, MuRF1, MuRF2, myomesin, NBR1, p62, T-cap) are depicted (structural proteins: purple; adapter proteins:
yellow; signaling proteins green). For the titin molecule the Ig domains (red), FN3 domains (white), Z-repeats
(green), the PEVK region (yellow), unique sequences (blue), and the kinase domain (black) are shown. Modified
from Bang et al., 2001a.
3.2.3.1 Z-disc region proteins
Titin’s N-terminus, which is integrated into the Z-disc of the sarcomere, forms cross bridges
with -actinin that links titin to the thin filament system (Ohtsuka et al., 1997a, b; Gregorio et
al., 1998). The protein sANK1 (small ankyrin 1) as well as T-cap (Titin-cap, telethonin, Figure
2) interact with titin’s Ig domains Z1 and Z2 (Kontrogianni-Konstantopoulos and Bloch, 2003;
Gregorio et al., 1998; Mues et al., 1998). The binding of sANK to titin suggests a role in
organizing the sarcoplasmic reticulum at the Z-disc of the sarcomere. T-cap together with its
binding partner MLP (muscle LIM protein) has been proposed to play a role as stretch sensor
(Knoll et al., 2002), in muscle growth (Nicholas et al., 2002), and in differentiation of myocytes
(Kong et al., 1997). Another binding partner of titin’s Z-disc region is obscurin that interacts
with titin’s Ig domains Z8 and Z9 (Bang et al., 2001a; Young et al., 2001). Obscurin plays a role
in signal transduction and co-assembles with titin during myofibrillogenesis (Sanger and Sanger,
2001; Young et al., 2001).
3.2.3.2 I-band region proteins
Within titin’s I-band region, the PEVK, N2A, and N2B region contain multiple protein binding
sites. FHL2 (four and a half LIM domain protein), which mediates protein-protein interactions,
binds to titin’s heart specific N2B region. FHL2 targets the metabolic enzymes
phosphofructokinase, creatine kinase, and adenylate kinase to titin to ensure energy supply at the
Introduction
12
site of high energy consumption (Lange et al., 2002). In addition to FHL, the small heat shock
protein B-crystallin interacts with the N2B region (Bullard et al., 2004). In response to
adrenergic stimulation, the N2B region can also be phosphorylated by the PKA (protein
kinase A) leading to the decrease of passive tension (Yamasaki et al., 2002). The N2A region
provides a binding site for the protease calpain 3 (Sorimachi et al., 1995), which is involved in
myofibrillogenesis, sarcomere assembly, and the proteolysis of different muscle proteins such as
titin, talin, and ezrin (Taveau et al., 2003; Kramerova et al., 2004). CARP (cardiac ankyrin
repeat protein), DARP (diabetes ankyrin repeat protein), and ANKRD2 (ankyrin repeat domain
protein) that are members of the MARP (muscle ankyrin repeat protein) protein family also bind
to titin’s N2A region (Bang et al., 2001b). They are involved in the modulation of gene
expression during development (Jeyaseelan et al., 1997; Baumeister et al., 1997; Bang et al.,
2001b). Another binding partner of titin’s N2A region is the protein myopalladin that influences
sarcomere integrity (Bang et al., 2001b). The PEVK region interacts with actin and the calcium-
binding protein S100A1 (S 100 calcium binding protein A 1). It has been speculated that this
titin-actin interaction contributes to muscle passive tension, which can be modulated by S100A1
(Yamasaki et al., 2001).
3.2.3.3 A-band region proteins
The FN3 and Ig domains that form the super-repeat structure within titin’s A-band region
interact with myosin and the myosin binding protein C (Figure 2), which also binds to myosin
(Okagaki et al., 1993), so that titin is linked to the thick filament (Wang et al., 1992; Houmeida
et al., 1995; Freiburg and Gautel, 1996). Theses interactions specify amount and location of
myosin and myosin binding protein C molecules that are incorporated into the thick filament
system. This it is believed to control the assembly and exact length of myosin filaments and to
act as a molecular scaffold maintaining the integrity of the sarcomere.
3.2.3.4 M-band region proteins
Titin’s C-terminal M-band region that is integrated into the M-band of the sarcomere provides at
its Ig domain M4 a binding site for the protein myomesin that forms antiparallel dimers
(Obermann et al., 1996). The additional interaction of myomesin with myosin and the direct
interaction of myosin with titin connect the titin molecules tightly to the thick filament to
compensate force imbalance in the sarcomere (Bähler et al., 1985; Nave et al., 1989; Obermann
et al., 1996). Furthermore titin’s M-band region binds MuRF1 (muscle-specific RING finger
Introduction
13
protein 1) as well as calpain 3 (Kinbara et al., 1997) and FHL2 (Lange et al., 2002), which both
also interact with titin’s I-band region (Gregorio et al., 1999). MuRF1 binds with its central
region to titin’s Ig domains A168 and A169 N-terminal to titin’s kinase region (Centner et al.,
2001). It has been shown that the interaction of MuRF1 with titin maintains the stability of the
filament’s structure (McElhinny et al., 2002). The binding sites for MuRF1 and calpain 3 from
opposing titin molecules are in close proximity. The functional relevance is unknown, but both
proteins have been implicated in ubiquitination and degradation processes (Bodine et al., 2001).
Cleavage of muscle proteins by the protease calpain 3 leads to ubiquitination and degradation
via the 26S proteasome (Hasselgren and Fischer, 2001). The ubiquitin ligase MuRF1 binds to
various muscle proteins including myosin light chain, T-cap, and nebulin suggesting a function
in controlling proteasome-dependent degradation (Witt et al., 2005). MuRF1 forms heterodimers
with the homologous proteins MuRF2 and MuRF3 (Centner et al., 2001; Gregorio et al., 2006),
which are involved in the stabilization of microtubules (Spencer et al., 2000; Pizon et al., 2002).
MuRF2 also interacts with titin’s domains A168-170 and other muscle proteins such as T-cap
and nebulin (Witt et al., 2005). MuRF1 interacts with enzymes that are involved in ATP-
production and with proteins that have nuclear function implicating a role in signal transduction
(Witt et al., 2005). Recent work shows that MuRF1 associates with SRF (serum response factor)
and interacts with RACK1 (receptor for activated C kinase 1) to regulate PKC (proteinase
kinase C) activity (Arya et al., 2004; Willis et al., 2007).
3.2.4 Titin’s kinase
Titin’s M-band region is encoded by six exons termed M-band exon 1-6, of which the M-band
exon 1 encodes a kinase domain (Labeit et al., 1992; Obermann et al., 1996). It is a
serine/threonine kinase that is highly conserved in vertebrates and invertebrates suggesting an
important function in signal transduction (Gautel et al., 1995). The kinase shows homology to
the myosin light chain kinase (MLCK) family and the invertebrate muscle proteins twitchin and
stretchin-MLCK (Heierhorst et al., 1994; Gautel et al., 1995). It consists of a catalytic domain
and a regulatory tail that blocks the ATP binding site. The activation of the kinase has been
proposed to base on a dual mechanism at the regulatory tail, in which the inhibiting tyrosine
residue Y170 is phosphorylated and Ca2+/calmodulin binds (Mayans et al., 1998). In contrast to
other kinases, which turn upon activation from a closed into an open conformation, titin’s kinase
already has the open conformation in the auto-inhibited form.
Introduction
14
The proteins T-cap, NBR1 (neighbor of Brca gene 1), and p62 (sequestosome 1) are suggested
substrates of titin’s kinase (Mayans et al., 1998; Lange et al., 2005). It has been shown that
titin’s kinase phosphorylates T-cap in differentiating cardiomyocytes indicating a role in
myofibrillogenesis (Mayans et al., 1998). Controversial, investigations in vivo have found that T-
cap expression starts after myofibrillogenesis at embryonic day 15.5 and that the loss of titin’s
kinase region does not affect sarcomere assembly until embryonic day 10.5 (Weinert et al.,
2006). Titin’s kinase has been implicated in responding to mechanical stress by force-induced
conformational changes (Grater et al., 2005). It has been shown that activation of the kinase by
stretch leads to a conformation that allows the interaction with NBR1, which targets the
ubiquitin-associated p62 to sarcomeres. The protein p62 interacts with MuRF2, which
translocates in response to mechanical inactivity to the nucleus, where it inhibits the
transcription activity of the SRF and reduces the amount of the SRF (Lange et al., 2005). A point
mutation in the human titin kinase domain causing myopathy was linked to this pathway (Lange
et al., 2005). Hence titin’s kinase has been implicated in the regulation of gene expression and
protein turnover. In contrast, results obtained with MuRF2-deficient mice suggest that titin-
MuRF2 signaling is dispensable for normal cardiac response to stress so that a molecular
mechanism by which titin’s kinase is involved in stress-sensing remains unclear (Willis et al.,
2007).
3.3 Smooth muscle titin
The first evidence for the existence of smooth muscle titin arose from the isolation of a protein
from chicken gizzard smooth muscle with an amino acid sequence similar to titin (Maruyama et
al., 1977). This protein has remained uncharacterized. Keller and colleagues have identified and
characterized a titin-like ~2000 kDa protein referred to as smitin (smooth muscle titin-like
protein) from chicken gizzard smooth muscle. They showed its localization within the smooth
muscle contractile apparatus as well as in vitro interactions with smooth muscle myosin (Keller
et al., 2000; Kim and Keller, 2002) and smooth muscle -actinin (Chi et al., 2005). Therefore it
is possible that smitin links dense bodies to thick filaments in avian smooth muscle. However,
smitin failed to cross-react with a tested titin antibody (Kim and Keller, 2002). Mahler and
colleagues have discovered in chicken the protein zeugmatin localizing at Z-discs of cardiac
muscle as well as dense bodies and attachment plaques of gizzard smooth muscle (Maher et al.,
1985). Later studies suggested that zeugmatin is the N-terminal region of titin (Turnacioglu et
al., 1996, 1997).
Introduction
15
The first investigation of titin in mammals revealed the expression of titin’s Z-disc region in
rabbit uterus using reverse transcription-PCR (RT-PCR; Sorimachi et al., 1997). Later it was
shown that human uterus, bladder, and carotid artery RNA contains titin transcripts encoding the
N-terminus including -actinin binding domains. Accordingly the binding of vertebrate smooth
muscle titin to smooth muscle -actinin in vitro has been demonstrated (Chi et al., 2008).
Furthermore, transcripts of titin’s exons 1 to 7 were amplified from the human smooth muscle
tissues aorta, stomach, carotids, and uterus by RT-PCR (Labeit et al., 2006). The expression of
titin isoforms in smooth muscle has been confirmed by a titin exon array, in which human aorta,
bladder, carotid, and stomach were investigated (Labeit et al., 2006). Alternatively spliced titin
isoforms were identified, which contain 80 to about 100 exons at a transcriptional level of ~100-
fold less abundant than in skeletal muscle. These exons encode parts of titin’s Z-disc, I-band,
and A-band regions. Consistent with the exon array data, a ~1000 kDa titin was detected by titin
Z-disc, I-band, and A-band antibodies in porcine aorta and stomach. Smooth muscle titin was
located in the cytoplasm of cultured human aortic smooth muscle cells and in the tunica media
of bovine aorta. As in avian smooth muscle, it has also been shown in vitro that vertebrate
smooth muscle titin contains binding sites for -actinin and filamins at its N-terminus and
binding sites for myosin at its C-terminus. Thus it has been suggested that smooth muscle titin
links thick filaments with dense bodies to provide passive elasticity and structural integrity to
smooth muscle.
3.4 Embryo spacing and implantation
3.4.1 The female reproductive system
The female reproductive system consists of vagina, uterus, oviduct, and ovaries. In the uterus
(corpus uteri) embryos implant and develop during pregnancy. It is a hollow organ located in the
pelvic cavity of female mammals. In higher primates the uterus is fused to a single organ, the
simplex form. In rodents such as mice the uterus consists of two separate uteri horns that are
dorsally combined, the duplex form (Figure 6). The uterus extends down into the cervix uteri
that leads into the vagina. On the top of the uterus two tubular oviducts, which are called
Fallopian tubes in woman, are connected. They link the uterus to the periovarian space and the
ovaries. In the duplex form, each uterine horn is associated with one oviduct.
The wall of the uterus is made up of the three layers endometrium (tunica mucosa), myometrium
(tunica muscularis), and perimetrium (tunica serosa). The endometrium lines the lumen of the
Introduction
16
uterus and consists of the lamina epithelialis mucosae and the lamina propria mucosae (stroma
endometrialis), which both undergo morphologic and functional changes in response to ovarian
hormones during the menstrual cycle in great apes and humans or estrous cycle in most other
mammals. The part of the endometrium that experiences changes (pars functionalis) is
reconstituted by the basal part (pars basalis; Hoffmeister and Schulz, 1961). The lamina
epithelialis mucosae is a single-layered prismatic epithelium with secretory and ciliated cells and
an underlying basal lamina. The lamina propria mucosae comprises of connective tissue that
encloses tubular glands (glandulae uterinae) and blood vessels. Next to the endometrium is the
myometrium that consists of an internal circular and an external longitudinal smooth muscle
layer, which are separated by highly vascularized connective tissue (stratum vasculosum). The
uterus is covered by a serous membrane, the perimetrium.
3.4.2 Embryo implantation
Mammalian embryo implantation ensures the firm connection of the embryo in the maternal
uterine tissue and involves the synchronizing of embryonic development until blastocyst state
with the uterine differentiation into the receptive state.
During ovulation unfertilized eggs (Figure 6 red) are released from the ovary into the
infundibulum of the oviduct. If mating occurs, the oocytes are fertilized leading to zygotes
(Figure 6 green). In mice, this time point is called embryonic day 0.5 (E0.5). The zygotes
undergo mitotic cell divisions leading to the formation of a morula until embryonic day 3.0
(E3.0). Thereby, the embryos travel along the oviduct by cilia transport that is amended by
smooth muscle contraction (Halbert et al., 1976, 1989; Vizza et al., 1995; Perez Martinez et al.,
2000). At embryonic day 3.5 (E3.5), embryos in the late morula stage consisting of totipotent
cells enter the uterine lumen and transform by lineage differentiation to an early blastocyst. They
are composed of a fluid filled cavity (blastocoel) as well as the trophectoderm and the inner cell
mass (ICM), which is pluripotent and generates the cell lineages of the embryo. Before
implantation, the blastocysts distribute equally among both uterine horns with evenly spacing so
that the implantation sites are equidistant independent of the blastocyst number. It has been
shown that this process of spacing involves smooth muscle contraction. Myometrial contractions
are higher during the preimplantation period and relaxin, an inhibitor of myometrial activity,
disrupts the blastocyst distribution in rats (Pusey et al., 1980; Rogers et al., 1983; Crane and
Martin, 1991). Recent findings indicate that lysophosphatic acid (LPA) signaling mediated by its
receptor LPA3 regulates embryo spacing by effecting uterine contraction (Hama et al., 2007).
Introduction
17
Figure 6: The murine reproductive system and preimplantational embryonic development. During ovulation
unfertilized eggs (red) are released into the oviduct, which are fertilized at embryonic day 0.5 (E0.5). Afterwards the
zygotes (green) develop thereby traveling along the oviduct to enter the uterine horns at embryonic day 3.5 (E3.5)
as blastocysts. Before implantation at embryonic day 4.0 (E4.0), the blastocysts are evenly distributed along both
uterine horns with evenly spacing (> E3.5). After embryonic day 4.0 (> E4.0), it is possible to make implantation
sites visible with the help of the dye Evans Blue (blue).
After hatching of the blastocysts from their outer shell (zona pellucida) and differentiation of the
ICM into the epiblast and primitive endoderm, implantation initiates. It is classified into
apposition, attachment, and penetration. During apposition at embryonic day 4.0 (E4.0), a fluid
accumulation in the intercellular tissue spaces of the uterus (edema) leads to uterine luminal
closure resulting in interaction of the embryonic trophectoderm with the luminal epithelium of
the maternal uteri endometrium. This is followed by the attachment phase at embryonic day 4.25
(E4.25), in which the trophectoderm associates tightly with the luminal epithelium. Apposition
and attachment depend on the synchronized differentiation and proliferation of the uterus to the
receptive state that occurs under the direction of the ovarian steroid hormones progesterone and
estrogen acting through their nuclear receptors (Carson et al., 2000; Dey et al., 2004). It has been
shown that the signaling pathway initiated by HB-EGF (heparin binding epidermal-like growth
factor) is crucial for blastocyst-uterine crosstalk (Paria et al., 1993; Raab et al., 1996). Growth
factors, adhesion molecules such as integrins, cytokines such as LIF (leukemia inhibitory
factor), and vasoactive mediators are also involved (Carson et al., 2000; Stewart et al., 1992;
Fouladi-Nashta et al., 2005). The attachment reaction involves a localized increase in stromal
vascular permeability of the uterus (Psychoyos, 1986), which is possible to visualize with the
help of the dye Evans Blue (Figure 6 blue). This vascular permeability is extended during the
penetration phase, when the embryo invades through the luminal epithelium and basal lamina
into the stroma. Thereby the superficial endometrial vessels are also penetrated by the embryo to
establish a vascular relationship (Schlafke and Enders, 1975) that requires uterine angiogenesis
as well (3.5.2). The stromal cells differentiate into decidual cells (decidualization) leading to the
loss of the luminal epithelium at the site of the implanting blastocyst (Parr and Parr, 1989). The
Introduction
18
deciduum provides nutritional support to the developing embryo before a functional placenta is
established.
3.5 Angiogenesis
During angiogenesis, new blood vessels develop from pre-existing vessels to ensure sufficient
blood transport, for example during pregnancy, wound healing, and inflammation. The newly
formed microvasculature consists of endothelial cells forming the inner vessel lining that is
covered by pericytes or vascular smooth muscle cells.
3.5.1 Sprouting angiogenesis
Angiogenesis by endothelial sprouting initiates with an increase in vascular permeability in
response to VEGF (vascular endothelial growth factor), a key regulator for vasculogenesis and
angiogenesis (Carmeliet et al., 1996; Fong et al., 1995). This allows leakage of plasma proteins
that provide a scaffold for migrating endothelial cells. Before endothelial cells migrate, their
interendothelial as well as the periendothelial cell contact are modulated to destabilize the
vessel. It has been shown that angiopoietin 2 is involved in detaching smooth muscle cells and
loosening the extracellular matrix (Maisonpierre et al., 1997; Gale and Yancopoulos, 1999).
Some endothelial cells within the capillary vessel are the tip cells that lead the growing sprout.
Notch receptors and their ligand DLL4 (delta-like-4) are essential for discriminating between tip
and the neighboring cells, which involves also VEGF (Sainson et al., 2005; Hellström et al.,
2007; Lobov et al., 2007; Suchting et al., 2007). The migration and proliferation of endothelial
tip cells is promoted by a spatial VEGF concentration gradient (Ruhrberg et al., 2002; Gerhardt
et al., 2003). The effects of VEGF are complemented by angiopoietin 1. This angiogenic factor
promotes remodeling including vessel maturation, stabilization, and leakiness by interaction
with the receptor Tie2 (tyrosine kinase with immunoglobulin-like and EGF-like domains 2; Suri
et al., 1996; Thurston et al., 1999; Sato et al., 1995; Wakui et al., 2006).
To form functional blood-carrying vessels, the endothelial sprouts need to suppress the motile
behavior and form endothelial-endothelial junctions with tip cells of other sprouts or with
existing capillaries. The generation of the vascular lumen stabilizes the new vessel by improved
oxygen delivery that lowers the Vegf expression. Vascular maturation requires also pericytes and
vascular smooth muscle cells. These mural cells are derived from neural crest cells,
undifferentiated mesenchymal cells, or vascular stem cells and have common molecular markers
Introduction
19
such as -smooth muscle actin, desmin, and the neuro-glial proteoglycan 2 (Armulik et al.,
2005; Bergers and Song, 2005). The endothelial sprout generates a concentration gradient of
PDGF-B (platelet derived growth factor B) that promotes the recruitment of mural cells (Lindahl
et al., 1997; Hellström et al., 1999). The tyrosine kinase receptor PDGF (PDGFR) that is
expressed in mesenchymal precursor cells, pericytes, and vascular smooth muscle cells mediates
the PDGF-B guided migration and is also required for mural cell proliferation and integration
into the vessel wall. PDGFR action involves cooperation with a family of G-protein coupled
sphingolipid receptors that bind the sphingolipid sphingosine-1-phosphate (S1P), which is
secreted by endothelial cells (Liu et al., 2000; Allende and Proia, 2002; Spiegel and Milstien,
2003). Mural cells that cover the endothelium of growing vessels inhibit endothelial cell
proliferation as well as migration and stimulate the formation of extracellular matrix proteins
thereby leading to vascular maturation (Gerhardt et al., 2003; Jain, 2003; Cleaver and Melton,
2003). Pericytes that are embedded in a basal lamina establish direct contact with endothelial
cells and cover immature blood vessels and capillaries. In contrast, vascular smooth muscle cells
are separated from the endothelium by a basement membrane layer consisting of fibers such as
collagen and cover mature and larger diameter vessels.
3.5.2 Uterine angiogenesis during embryo implantation
Angiogenesis takes place in the ovary and uterus of adult mammals during the reproductive
cycle and pregnancy (Gordon et al., 1995; Torry and Torry, 1997; Abulafia and Sherer, 1999). In
the preimplantation period, increased uterine permeability as well as angiogenesis occurs at the
site of blastocyst apposition to provide sufficient vascularized uterine tissue for successful
implantation (Plaks et al., 2006). A number of studies show that VEGF and its receptors
VEGFR1, VEGFR2, and NRP1 (neuropilin 1) are differentially expressed in mouse uterus in a
spatiotemporal manner before and during the attachment phase (Hyder and Stancel, 1999;
Halder et al., 2000; Chakraborty et al., 1995). Furthermore, it has been shown in uterine samples
of women with recurrent miscarriage that the expression of the receptors for VEGF and
angiopoietins was reduced (Vuorela et al., 2000; Vuorela and Halmesmäki, 2006). Inhibition of
angiogenesis before or after implantation by an antiangiogenic compound results in resorption of
all embryos in mice (Klauber et al., 1997) suggesting that the formation of new blood vessels
from pre-existing vessels is important for implantation and development of the placenta.
During the preimplantation period, Vegf expression and uterine vascular permeability, which is
required for angiogenesis, occurs in mouse in response to the ovarian steroid hormones estrogen
Introduction
20
and progesterone (Chakraborty et al., 1995; Hyder et al., 2000; Ma et al., 2001). Furthermore, it
has been shown that VEGF is mainly responsible for the estrogen-induced increase in vascular
permeability, which is essential for successful embryo implantation (Rockwell et al., 2002).
Results obtained from mice carrying a null mutation for Cox2, the gene encoding the rate-
limiting enzyme in prostaglandin biosynthesis cyclooxygenase 2 (COX2), indicate that
prostaglandins influence uterine vascular permeability and angiogenesis during implantation and
decidualization via VEGF and angiopoietin signaling (Matsumoto et al., 2002). It was shown
that HB-EGF produced by embryo and uterus induces Cox2 expression in the luminal epithelium
and underlying stromal cells at the site of blastocyst attachment (Lim et al., 1997).
3.6 Knockout and knockin technology
Knockout and knockin mice are genetically engineered so that a gene is turned off or modified
by an insertion, respectively. The technology is based on the usage of pluripotent embryonic
stem (ES) cells as first described by Evans and Kaufmann (Evans and Kaufman, 1981).
Differentiation of the ES cells in cell culture is inhibited to allow contribution to germline cells
of chimeras. Therefore, ES cells are grown on mitotic inactivated primary embryonic fibroblasts
(feeder cells) and the protein LIF is added to the culture medium (Smith et al., 1988).
To modify the gene of interest by homologous recombination, a recombinant targeting vector is
cloned in vitro, which contains a long and a short arm as well as a middle part. The long and the
short arm consist of DNA sequences that are homologous to the gene of interest. The middle part
contains a neomycin resistance cassette flanked by two FRT-sites (flippase recognition target-
sites) and the modified gene. The targeting vector enters ES cells by electroporation, in which a
short current pulse opens the cell membrane (Piedrahita et al., 1992; Austin et al., 2004), and
integrates in a few cells stably in the genome. The integration occurs either randomly or into the
gene locus of interest by homologous recombination due to homologous sequences of the
targeting vector (Doetschman et al., 1987; Thomas and Capecchi, 1987). The antibiotic G418
that inhibits protein translation is added to the cell medium to select the ES cells, which
integrated the targeting vector stably into the genome. Only these cells contain the neomycin
resistance cassette encoding the bacteria gene for the neomycin phosphotransferase under the
control of a ubiquitous promoter. This enzyme degrades the antibiotic so that the ES cells can
grow in the presence of G418. Antibiotic-resistant cells are isolated and investigated by PCR or
Southern Blot for homologous recombination of the targeting vector into the genome (Piedrahita
et al., 1992; Austin et al., 2004). Homologously recombined ES cells are injected into
Introduction
21
blastocysts that are obtained from superovulated female mice. The injected blastocysts are
transferred into the uteri of pseudopregnant mice, in which they implant so that embryos can
develop. Offspring of the recipient mice are called chimeras because they are arisen from cell
populations of both blastocyst and recombined ES cells. In contrast to ES cells, which have been
generated from a mouse strain with a brown (agouti) coat color (129), the blastocysts have been
obtained from a mouse strain with a black coat color (C57BL/6) so that chimeric animals show a
black and brown coat color mixture (Bradley et al., 1984). Chimeras are bred with the mouse
strain C57BL/6. This breeding results in brown offspring if germline cells of the chimeras have
been developed from the heterozygously recombined ES cells. Half of these animals should be
heterozygous for the modified gene.
Since the neomycin resistance cassette has the potential to affect the phenotype of genetically
engineered animals (Kaul et al., 2000), heterozygous animals carrying the targeting vector are
mated to transgenic mice that express the Flp-recombinase in their germline (Rodriguez et al.,
2000). This enzyme catalyzes the recombination between two FRT-sites thereby cutting out any
DNA stretch between. Offspring from this mating, in which the neomycin cassette has been
removed by Flp-mediated excision, is used to generate a colony of homozygous mice that only
contain the modified gene and one residual FRT-site.
3.7 Aim of the study
Human infertility is a global problem. ~12% of women have difficulties in getting pregnant or
carrying a baby to term (NSFG, 2002). Today conception can be improved by in vitro
fertilization and embryo transfer techniques. Nevertheless, implantation failure and pregnancy
loss are not only common problems after assisted reproduction, but also after natural conception.
Furthermore, fetal as well as maternal health is endangered by ectopic pregnancies as well as
placenta previa, which are associated with mislocalization of the embryo. The mechanisms of
smooth muscle contraction that contribute to the localization of the embryo within the uterus are
not well understood. Cellular events of implantation have been described, but only little is
known about the molecular pathways. A hallmark of implantation is uterine vascular
development and remodeling at the fetal-maternal interface triggered by angiogenic factors. The
importance of vessel formation has been demonstrated because disruption of angiogenesis at the
site of implantation is associated with poor reproduction in humans (Meegdes et al., 1988;
Vuorela et al., 2000). It is necessary to define the mechanism by which defects in angiogenesis
Introduction
22
contribute to human implantation failure and early miscarriage. This will help to improve fetal
health and female fertility.
Smooth muscle cells are involved in angiogenesis and responsible for uterine contraction. In
contrast to striated muscle, in which sarcomeres have been well characterized, structure and
signaling of the smooth muscle contractile unit need to be investigated in more detail. The actin-
myosin based structural and functional analogy between the smooth and striated muscle
contractile apparatus suggests that smooth muscle also contains a titin-like protein involved in
structure, biomechanics, and signal transduction. Although smooth muscle titin has been
described on transcript and protein level, it is not included in the current models of smooth
muscle structure and a function has not been assigned. Aim of our research was the
morphological and functional characterization of titin in smooth muscle. A knockin mouse
model should be established to visualize and thereby localize titin in situ and in vivo using the
red fluorescence protein DsRed. To address a role for smooth muscle titin, we used a loss of
function approach of titin’s kinase region. We hypothesized that titin’s kinase region in smooth
muscle has essential functions regarding muscle contractility and signaling as it has been shown
for embryonic as well as mature striated muscle (Gotthardt et al., 2003; Weinert et al., 2006;
Peng et al., 2006, 2007). Furthermore, it is possible that titin’s kinase in smooth muscle converts
mechanical input to biochemical signals by interacting with signal transduction proteins in a
strain-dependent manner similar to its proposed function in striated muscle (Tskhovrebova and
Trinick, 2003; Granzier and Labeit, 2004; Lange et al., 2005). The clinical relevance of titin’s
M-band region has been demonstrated by a point mutation in the human titin kinase domain that
causes myopathy (Lange et al., 2005). For our study, the conditional knockout model of titin’s
M-band exons 1 and 2 (Gotthardt et al., 2003), which encode titin’s kinase region, was available
to generate a smooth muscle knockout mouse strain. We used this mouse model to assess,
whether the deletion of titin’s kinase region affects embryo spacing and implantation as well as
angiogenesis. Furthermore, insights into signaling pathways involving titin’s kinase region
during early pregnancy should be gained with the help of Real-Time PCR, 2D-gel analysis, and
immunoassays.
Materials and Methods
23
4 Materials and Methods
4.1 Materials
4.1.1 Chemicals
If not stated otherwise, all chemicals were purchased from Sigma-Aldrich, Roth, Invitrogen, and
GE Healthcare.
4.1.2 Enzymes
Table 1: Enzymes.
Name Manufacturer
Calf intestinal alkaline phosphatase (CIP) Roche
Collagenase type I Sigma-Aldrich
Collagenase type II Invitrogen
Dispase II Sigma-Aldrich
Elastase type IV Sigma-Aldrich
Expand Long Template PCR System Roche
Phusion High-Fidelity DNA Polymerase New England BioLabs
Proteinase K Roche
Restriction endonucleases New England BioLabs and MBI Fermentas
RNase A from bovine pancreas (type III-A) Sigma-Aldrich
Taq DNA polymerase Invitrogen
4.1.3 Bacterial strain
For transformation of plasmid DNA, the Escherichia coli (E. coli) strain DH5 (Bethesda
Research Laboratories, 1986) was used.
Materials and Methods
24
4.1.4 Vectors and BAC clones
For cloning of the targeting vector, the plasmids and bacterial artificial chromosome (BAC)
clones listed in Table 2 were used.
Table 2: Plasmids and BAC clones used for generation of the targeting vector.
Name Manufacturer
BAC clone 96022 CITB Mouse BAC Library, Incyte Genomics
BAC clone RP23-436E5 BACPAC Resource Center
pDsRed-Monomer-C1 Clontech
pGemFRTloxNeo Prof. Dr. med. Gotthardt
4.1.5 Kits
Table 3: Kits.
Name Manufacturer
12(S)-HETE EIA Kit Assay Designs
Agilent RNA 6000 Nano Kit Agilent
BigDye Terminator v3.1 Cycle Sequencing Kit Applied Biosystems
DNA Ligation Kit Ver.1 TAKARA BIO INC.
EasyPure DNA Purification Kit Biozym
FAST qPCR MasterMix Plus Eurogentec
Growth Factor Mouse 4-Plex Panel Invitrogen
MILLIPLEX Mouse Cytokine/Chemokine Panel-7-Plex Millipore
NucleoBond BAC 100 Macherey-Nagel
peqGOLD Plasmid Miniprep Kit I PEQLAB
pGEM-T Easy Vector System Promega
Plasmid Maxi Kit Qiagen
RNase-Free DNase Set Qiagen
RNA UltraSense One-Step Quantitative RT-PCR System Invitrogen
RNeasy Mini Kit Qiagen
ThermoScript RT-PCR System for First-Strand Synthesis Invitrogen
4.1.6 Oligonucleotides
Primers for cloning, sequencing, and genotyping (Table 4) were designed using the online
software Primer3. Oligonucleotides for cloning were synthesized by TIB MOLBIOL and
oligonucleotides for sequencing and genotyping were synthesized by BioTeZ. Primer and probes
Materials and Methods
25
for Real-Time PCR analysis were purchased from Applied Biosystems as gene expression assays
(Alox15, Cacna1c, Calb1, Calm1, Camk2a, Cox2, Creb1, Fhl2, Lpar3, Mapk13, Myom1, Nbr1,
p62, Pparg, Prkcd, Prkce, S100a1, T-cap) or designed using the Software Primer Express 1.5
(Applied Biosystems) to be synthesized by BioTeZ or Eurogentec. The double labeled probes for
Real-Time PCR analysis carried the fluorescence reporter dye 6-carboxyfluorescein (6-FAM) at
the 5’-end and the quencher dye 6-carboxytetramethylrhodamine (TAMRA) at the 3’-end (Table
5). The sequences of the oligonucleotides that were custom made are provided in
5’3’orientation in the following tables.
Table 4: Primers for cloning, sequencing, and genotyping provided in 5’3’orientation.
Name Sequence Application
3’neoflox TCGACTAGAGGATCAGCTTGGGCTG genotyping
Cre800fw GCTGCCACGACCAAGTGACAGCAATG genotyping
Cre1200rev GTAGTTATTCGGATCATCAGCTACAC genotyping
lacZ1080fw CCTCTGCATGGTCAGGTCATGGATG genotyping
lacZ10620rev GTGGGCGTATTCGCAAAGGATCAGC genotyping
MG-FLP1 GTCACTGCAGTTTAAATACAAGACG genotyping
MG-FLP2 GTTGCGCTAAAGAAGTATATGTGCC genotyping
MG-Ti-FRTr2 AAGTTCGCTATACAACTGAGGCTAAG genotyping
MG-Ti-SL1fw GTGTCTGGCACTGCTTCCTTGGAAGTG genotyping
MG-Ti-SL2rev ACCGCTCCCATGCCTTCGAGAGTCTTG genotyping
MR-GK2Prom5’r AAAGCGCATGCTCCAGACTGCCTTG sequencing
MR-neor AGCCATGATGGATACTTTCTCG sequencing
NB-DsRedmf CTCCACCGAGAAGCTGTACC sequencing
NB-DsRedmr CTTGGAGCCGTACTGGAACT sequencing
NB-fDsRedI GGTACCCTCGAGATGGACAACACCGAGGACGTCA cloning
NB-fDsRedII CTCGAGATGGACAACACCGAGGACGTCA genotyping
NB-fDsRedrecF CAGCATCATGGTAAAGGCCATCAA genotyping
NB-fLA CTCGAGAGTTCCCCACACTTTGAAGAAAG cloning
NB-fMA TGTACAGTGAAGCTGAAAAGGGCTGAAAGG cloning
NB-fSA GGATCCATATTTAAATGTTTCTCTTTACCTATTATTCAAG cloning
NB-GKFRTr GGGGGAACTTCCTGACTAG sequencing
NB-LOXFRTf CTGGGGCTCGACTAGAGGAT sequencing
NB-rDsRedI GGTACCCTGGGAGCCGGAGTGGCG cloning
NB-rDsRedII CTGGGAGCCGGAGTGGCG genotyping
NB-DsRedGeno ATAGAGACCTTGCTTTGCCTGTG genotyping
NB-rDsRedrecF CATTCAAATGTTGCCATGGTGTCC genotyping
Materials and Methods
26
NB-rLA CTCGAGCACTTCTTTGGTTTCGCTGGT cloning
NB-rMA AAGCTTGATAAGGGATAGTCTTGGGCATAC cloning
NB-rSA CCCGGGTCACTCTTTTCAGACACACCAGA cloning
NB-rSAc CCCGGGCTGCAAAAGAAGACACACAGAAAG cloning
NB-ttnEx18f GGGTATTTTGTGCCAAGCTC sequencing
NB-ttnEx19r TCCTCTGTGATGCTGGTGTC sequencing
NB-ttnEx20f CCACCGAAGAGAAACGGTAA sequencing
NB-ttnEx21f GAGCTTAGATGTGGGCCGTA sequencing
NB-ttnEx22f TTTGGAAAATCTGGGTTTGG sequencing
NB-ttnEx23f TCAATAAGCCTATAACACTGGAACT sequencing
NB-ttnEx24r ATTTGCTCTTGGGATGTTCG sequencing
NB-ttnEx25f TCATTCTAAAACATGCAGAAGCA sequencing
NB-ttnEx26f CCATGAGTCCCTGAACACAG sequencing
NB-ttnEx27f GGTAAAAACAAGAGTTTTATTGAAATG sequencing
NB-ttnEx27r TGGTGGTAGCACTTTCTGCTT sequencing
NB-ttnEx28f TACCCAAGGATCAGGCATGT sequencing
NB-ttnEx28mf CAACTGCCTGATGGGAAAAA sequencing
NB-ttnEx28mr GCTGAGGGTAGCCTGTCACT sequencing
NB-ttnEx30mr TCCCATCAACAACAAAGCTG sequencing
Sp6 GGATCCCTATACTTCAGAGTCTTCT sequencing
T7 TAATACGACTCACTATAGG sequencing
Table 5: Primer/probes for Real-Time PCR analysis provided in 5’3’orientation (fw: forward; rev: reverse;
FAM: 6-carboxyfluorescein; TAMRA: 6-carboxytetramethylrhodamine).
Name Sequence
18S RNA fw CGCCGCTAGAGGTGAAATTC
18S RNA rev TGGGCAAATGCTTTCGCTC
18S RNA probe 6-FAM-TGGACCGGCGCAAGACGGAC-TAMRA
MEx1-2 fw CCGATGGACTCAAGTACAGGATT
MEx1-2 rev CCCATGCCTTCGAGAGTCTT
MEx1-2 probe 6-FAM-TCCTTGGAAGTGGAAGTTCCAGCTAAGATACAC-TAMRA
MEx6 fw GCCTTGTGTGGTAGTTCTAAATTCAA
MEx6 rev TTTGCTGTGGCTCATTGCTT
MEx6 probe 6-FAM-TTTCACCGGGAACTGGGCAA-TAMRA
MuRF1 fw CCGAGTGCAGACGATCATCTC
MuRF1 rev CCTTCACCTGGTGGCTATTCTC
MuRF1 probe 6-FAM-AGCTGGAGGACTCGTGCAGAGTGACC-TAMRA
Materials and Methods
27
MuRF2 fw TGGAGAACGTATCCAAGTTGGT
MuRF2 rev CCTTTGATGCTTCCACGATCT
MuRF2 probe 6-FAM-CATGGATGAGCCCGAAATGGCA-TAMRA
N2A fw AAAATGTGGATCCTAAAGAGTATGAGAAG
N2A rev GAGGAGGCCCCGGAAGT
N2A probe 6-FAM-TGCGCGCATGTACGGAATCACC-TAMRA
ZEx3 fw CGATGGCCGCGCTAGA
ZEx3 rev CTCAGGGAGTATCGTCCACTGTT
ZEx3 probe 6-FAM-TGATGATCCCCGCCGTGACTAAAGC-TAMRA
4.1.7 Antibodies
Primary and secondary antibodies, which were used for immunofluorescence staining and
Western blotting, including the host species, dilutions, and manufacturer are provided in the
tables below.
Table 6: Primary antibodies used for immunofluorescence staining (IF) and Western blotting (WB).
Name Species Dilution IF Dilution WB Manufacturer
-actinin (EA53) mouse 1:500 Sigma-Aldrich
DsRed rabbit 1:200 1:500 Clontech
smooth muscle actin rabbit 1:100 Lab Vision
titin M8/M9 rabbit 1:200 1:500 Prof. Dr. Siegfried Labeit
Table 7: Secondary antibodies used for immunofluorescence staining (IF) and Western blotting (WB).
Name Species Dilution IF Dilution WB
Manufacturer
anti-mouse Alexa Fluor 488 goat 1:500 Invitrogen
anti-rabbit Cy3 goat 1:500 Jackson Research
anti-rabbit IgG HRP-conjugated goat 1:2000 GE Healthcare
anti-rabbit IgG biotin-conjugated goat 1:500 Jackson Research
Materials and Methods
28
4.2 Methods
4.2.1 Molecular biology methods
4.2.1.1 DNA preparation
4.2.1.1.1 Preparation of plasmid DNA
Bacteria colonies generated by transformation (4.2.2.2) were used to inoculate overnight
cultures. Small amounts of recombinant plasmid DNA for cloning were isolated from 3 ml
cultures with the peqGOLD Plasmid Miniprep Kit I. Higher yield of plasmid DNA of the initial
cloning vectors pDsRed-Monomer-C1 and pGemFRTloxNeo as well as the targeting vector
DsRedKI was obtained by processing 200 ml cultures using the Plasmid Maxi Kit. Both kits
were used according to the protocol given by the companies. The DNA concentration was
determined (4.2.1.2) and the DNA was stored at -20°C.
4.2.1.1.2 Preparation of BAC DNA
BACs were isolated from 500 ml overnight cultures of the BAC clones 96022 and RP23-436E5
using the NucleoBond BAC 100 Kit according to the “Low-copy plasmid purification
(Maxi/BAC, Mega)” protocol provided by the manufacturer. Before storage at -20°C, the
concentration was determined (4.2.1.2).
4.2.1.1.3 Purification of DNA from agarose gels
For cloning, DNA fragments were extracted from agarose gels (4.2.1.4) using the EasyPure
DNA Purification Kit according to manufacturer’s instructions. DNA was eluted in 15 l double
distilled water (ddH2O). Afterwards, the concentration was measured and storage was carried
out at -20°C.
4.2.1.1.4 Preparation of genomic DNA from ES cells
96-well cell culture plates, in which ES cells were grown to confluence, were obtained from the
MDC Transgenic Core Facility (4.2.3.1) and used for preparation of genomic DNA. Cell culture
Materials and Methods
29
medium was removed and the ES cells were washed twice with 100 l phosphate buffered saline
(PBS; 137 mM NaCl, 2.7 mM KCl, 4.3 mM Na2HPO4×2 H2O, 1.4 mM KH2PO4, pH 7.4). 100 l
ES cell lysis buffer (0.1% sodium dodecyl sulfate (SDS)) supplemented with 2 µl proteinase K
(14 mg/ml) and 0.5 l RNase A (10 mg/ml) was added to each well for digestion of the cells
overnight at 55°C in a wet chamber. DNA was precipitated overnight using 10 l 8 M LiCl and
100 l isopropanol and subsequent centrifugation for 30 min at 3500 rpm and 4°C. DNA pellets
were washed with 70% ethanol, air-dried and dissolved in 50 l TE (Tris-EDTA) buffer (10 mM
Tris-HCl, 1 mM Na2EDTA, pH 8.0) at 55°C in a wet chamber overnight. After measuring of the
concentration (4.2.1.2), the DNA was stored at 4°C.
4.2.1.1.5 Preparation of genomic DNA from mouse tail
For genotyping with PCR products exceeding 600 bp, genomic DNA was purified from the tail
tip. Therefore, ~2 mm of the tail tip (4.2.5.1) was digested in 725 l tail buffer (20 mM Tris-base
pH 8.0, 5 mM Na2EDTA pH 8.0, 0.2% SDS, 4.3 mM NaCl) supplemented with 25l
proteinase K (10 mg/ml) at 52°C with 750 rpm shaking overnight. DNA was extracted with
phenol/chloroform/isoamyl alcohol (25:24:1) and precipitated by the addition of 0.1 volume 8 M
LiCl and 2.5 volumes 99% ethanol. After centrifugation at 13,000 rpm and 4°C for 30 min, the
pellet was washed twice with 70% ethanol, air-dried and resuspended in 100 l TE buffer
(4.2.1.1.4). The concentration was determined (4.2.1.2) and samples were stored at 4°C.
4.2.1.1.6 Preparation of short DNA fragments from mouse tissue
Mouse genomic DNA for PCR genotyping was prepared following the HotSHOT method
(Truett et al., 2000), if the expected PCR fragments had a size of less than 600 bp. ~1 mm of
mouse tail snips (4.2.5.1) or uterine tissue in estrous stage (4.2.5.7) were heated at 95°C for
30 min in 75 l alkali lysis buffer (25 mM NaOH, 0.2 mM Na2EDTA). After cooling on ice,
75 l neutralization buffer (40 mM Tris-HCl) was added and 1 l was directly used for PCR.
DNA preparations were stored at 4°C for short term or at -20°C for long term.
4.2.1.2 Determination of nucleic acid concentration
The concentration of DNA and RNA samples were measured spectrophotometrically at 260 nm
using the NanoDrop ND-1000 (Thermo Scientific) according to manufacturer’s instructions.
Materials and Methods
30
4.2.1.3 Polymerase chain reaction (PCR)
All polymerase chain reactions (PCRs) were carried out in the Hybaid Px2 Thermal Cycler
(Thermo Scientific) or in the PTC-200 and PTC-225 Thermo Cycler (MJ Research) using
primers as listed in Table 4 and dNTPs from Invitrogen or a Kit for 4.2.1.3.1 and 4.2.1.3.2.
4.2.1.3.1 Amplification of DNA for cloning
DNA fragments for cloning were amplified from plasmids or BAC clones. Table 8 shows the
primer pairs, the template, and the expected product size for each PCR.
Table 8: Primers, template, and expected sizes of the PCR products for cloning.
PCR Primer Template Product Size
DsRed NB-fDsRedI, NB-rDsRedI 5 ng pDsRed-Monomer-C1 693 bp
long arm (LA) NB-fLA, NB-rLA 1 ng BAC clone RP23-436E5 7838 bp
middle arm (MA) NB-fMA, NB-rMA 1 ng BAC clone 96022 654 bp
short arm (SA) NB-fSA, NB-rSA 1 ng BAC clone 96022 1592 bp
short arm control (SAc) NB-fSA, NB-rSAc 1 ng BAC clone 96022 1871 bp
The PCRs DsRed, middle arm, short arm, and short arm control were performed using the
Expand Long Template PCR System preparing the reaction mix according to manufacturer’s
instructions with 1 or 5 ng template (Table 8). The PCR products were amplified using the
following cycle conditions, whereas the elongation time was for the DsRed and MA PCR 45 s
and for the SA and SAc PCR 75 s:
1. initial denaturation 94°C 120 s
2. denaturation 94°C 15 s
3. annealing 55°C 30 s
4. elongation 72°C 45 or 75 s 10 x step 2-4
5. denaturation 60°C 45 s
6. annealing 65°C 15 s
7. elongation 65°C 45 or 75 s + 5 s/cycle 20 x step 4-6
8. final elongation 72°C 7 min
9. storage 4°C ∞
Materials and Methods
31
The LA PCR to amplify the long arm was done with the Phusion High-Fidelity DNA
Polymerase utilizing the pipetting instruction as given by the company with 1 ng of template
(Table 8). The PCR reaction was done according to the program:
1. initial denaturation 98°C 30 s
2. denaturation 98°C 10 s
3. annealing 60°C 30 s
4. elongation 72°C 150 s 29 x step 2-4
5. final elongation 72°C 10 min
6. storage 4°C ∞
After PCR, DNA fragments were loaded on an agarose gel (4.2.1.4), purified (4.2.1.1.3) and
subcloned into the pGEM-T Easy Vector (4.2.1.6.2.4). Before subcloning, the LA PCR product
was A-tailed (4.2.1.6.2.1).
4.2.1.3.2 DNA sequencing
The PCR products used for cloning and the targeting vector DsRedKI were verified by DNA
sequencing using the chain termination method. The BigDye Terminator v3.1 Cycle Sequencing
Kit was utilized for the sequencing reactions with the primers as listed in Table 4 according to a
protocol modified from manufacturer’s instructions. The PCR fragments short arm, DsRed,
middle arm, long arm, and short arm control inserted into the pGEM-T Easy vector were
sequenced with the primers Sp6 and T7, whereas the long arm was additionally sequenced with
primers covering titin’s exons 18 to 28 (Table 4). The complete targeting vector DsRedKI was
sequenced with all primers for sequencing that are listed in Table 4, except Sp6. As template,
300 ng of purified plasmid DNA (4.2.1.1.1) was used in the following reaction mixture:
final concentration volume
5 x BigDye sequencing buffer 1 x
2 l
10 M primer 250 nM
0.25 l
DNA template 300 ng/10 l
X l
2.5 x BigDye Ready Reaction Premix 0.25 x
1 l
ddH2O
add to 10 l
Materials and Methods
32
The PCR cycling conditions were chosen:
1. denaturation 96°C 10 s
2. annealing 50°C 5 s
3. elongation 60°C 4 min 30 x step 1-3
4. storage 4°C ∞
PCR products were purified by gel filtration using Sephadex G50 superfine (GE Healthcare) in
MultiScreen Filter Plates (Millipore) following the guidelines of the company. 10 l Hi-Di
Formamide (Applied Biosystems) was added to each purified sample. The sequencing was done
with a 3100 Genetic Analyzer (Applied Biosystems) that was operated by the group of Professor
Dr. Norbert Hübner. The sequence data were analyzed with the help of the software
SeqMan II 4.03 (DNASTAR Inc.).
4.2.1.3.3 Soriano PCR for Es cell and mouse genotyping
An adapted Soriano protocol (Nagy et al., 2003a) was used for the neo PCR to screen ES cells
for homologous recombination of the targeting vector and to verify the integration of the
targeting vector into the titin locus of chimeras and their offspring. 10 x Gittschier buffer
(166 mM (NH4)2SO4, 670 mM Tris-HCl, 67 mM MgCl2, 50 mM -mercaptoethanol, 67 M
Na2EDTA, pH 8.8), the primers 3’neoflox and NB-DsRedGeno, and 300 ng ES cell DNA
(4.2.1.1.4) or 100 ng mouse genomic DNA (4.2.1.1.5) as template were utilized leading to a
PCR product with a size of 1971 bp. The following PCR reaction mix was used:
final concentration volume
10 x Gittschier buffer 1 x
2.5 l
DMSO 10 %
2.5 l
10 mg/ml BSA 80 g/ml
0.2 l
10 mM dNTP mix 1 mM
2.5 l
10 M primer forward 0.4 M
1 l
10 M primer reverse 0.4 M
1 l
DNA template 100 or 300 ng/25 l
X l
5 U/l Taq DNA polymerase 0.05 U/l
0.25 l
ddH2O
add to 25 l
The PCR cycle profile was:
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1. initial denaturation 94°C 80 s
2. initial annealing 58°C 120 s
3. initial elongation 65°C 5 min
4. denaturation 94°C 30 s
5. annealing 58°C 30 s
6. elongation 65°C 180 s 40 x step 4-6
7. final elongation 65°C 10 min
8. storage 10°C ∞
The PCR reactions were loaded on an agarose gel for detection of the products (4.2.1.4).
4.2.1.3.4 PCR for mouse genotyping
For mouse genotyping the PCRs as listed in Table 9 were used.
Table 9: Primers, PCR conditions, and expected product sizes of the genotyping PCRs.
PCR Primer MgCl2 Annealing Product Size
cre Cre800fw, Cre1200rev 1.5 mM 55°C 400 bp
DsRed NB-fDsRedII, NB-rDsRedII 1.5 mM 57°C 693 bp
FLP MG-FLP1, MG-FLP2 3 mM 55°C 480 bp
lacZ lacZ1080fw, lacZ10620rev 1.5 mM 55°C 550 bp
lox MG-Ti-SL1fw, MG-Ti-SL2rev 3 mM 55°C WT 200 bp/ lox 300 bp
rec MG-Ti-SL1fw, MG-Ti-FRTr2 2 mM 55°C 370 bp
recF NB-fDsRedrecF, NB-rDsRedrecF 1.5 mM 57°C WT 282 bp/ recF 369 bp
The following protocol was used with 1 l of HotSHOT DNA (4.2.1.1.6) and final MgCl2
concentrations as indicated in Table 9:
final concentration volume
10 x Taq DNA polymerase Buffer 1 x
2.5 l
50 mM MgCl2 1.5 or 2 or 3 mM
0.75 or 1 or 1.5 l
10 mM dNTP mix 0.2 mM
0.5 l
10 M primer forward 0.4 M
1 l
10 M primer reverse 0.4 M
1 l
DNA template
1 l
5 U/l Taq DNA polymerase 0.03 U/l
0.15 l
ddH2O
add to 25 l
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The following PCR amplification conditions were chosen with annealing temperatures as given
in Table 9:
1. initial denaturation 94°C 120 s
2. denaturation 94°C 15 s
3. annealing 55 or 57°C 45 s
4. elongation 72°C 180 s 35 x step 2-4
5. final elongation 72°C 8 min
6. storage 10°C ∞
The PCR reactions were analyzed using agarose gels (4.2.1.4).
4.2.1.4 DNA agarose gel electrophoresis
Size dependent separation and visualization of DNA fragments after PCR and restriction digest
was done using agarose gels. Depending on the expected DNA fragment sizes, 0.5-2% agarose
gels were prepared using UltraPure Agarose (Invitrogen) in 0.5 x Tris-acetate-EDTA (TAE)
buffer (20 mM Tris-HCl, 5 mM Na-acetate, 0.5 mM Na2EDTA, pH 8.0) supplemented with
ethidium bromide solution to a final concentration of 0.5 g/ml. Samples were mixed with 0.1
volume of DNA loading buffer (50% glycerol, 0.5% orange G, 25 mM Na2EDTA) and loaded on
a gel to separate DNA fragments in 0.5% TAE buffer using a Mupid-21 Mini-Gel
Electrophoresis Unit (Cosmo Bio Co.) at 100 V. DNA fragments were visualized at 312 nm and
photographed with the Gel Doc 2000 Gel Documentation System (Bio-Rad) and their size was
determined using the 1 Kb DNA Ladder (Invitrogen).
DNA fragments, which were used for cloning of the targeting vector, were cut out with a scalpel
under UV light and purified (4.2.1.1.3).
4.2.1.5 Real-Time PCR
4.2.1.5.1 Preparation of RNA
Tissue of non-pregnant (estrous stage) and E3.5 pregnant SMcTiMEx1-2 mice (4.2.5.7) was
transferred into 1 ml Trizol (Invitrogen) to isolate total RNA according to the manufacturer’s
recommendations. The tissue was homogenized 3 x 20 s using an Ultra-Turrax T-25 (IKA-Werk)
followed by the optional centrifugation step. Isolated RNA was dissolved in 89 l RNase-free
water (Qiagen) and stored at -80°C.
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Isolated total RNA was thawed on ice and the RNA concentration was determined (4.2.1.2).
100 g of RNA were digested with the RNase-Free DNase Set to remove genomic DNA
according to the protocol “DNase Digestion of RNA before RNA Cleanup” of the Qiagen
RNeasy Mini Handbook. Afterwards the RNA was purified using the RNeasy Mini Kit
according to manufacturer’s protocol for RNA Cleanup. RNA was eluted with 2 x 30 l of
RNase-free water. The concentration of the RNA was again determined (4.2.1.2) and the RNA
quality was assessed using the Bioanalyzer (4.2.1.5.2).
4.2.1.5.2 Bioanalyzer
RNA quality was determined using the 2100 Bioanalyzer (Agilent). This method is based upon
on-chip gel electrophoresis and detection by laser-induced fluorescence of dye labeled RNA.
RNA was processed using the Agilent RNA 6000 Nano Kit together with the Agilent RNA 6000
Ladder following the given protocol. Only samples with a RNA integrity number (RIN) of over
8 were used for cDNA synthesis (4.2.1.5.3) or Real-Time PCR analysis (4.2.1.5.4).
4.2.1.5.3 cDNA synthesis
To convert RNA into complementary DNA (cDNA), a reversed transcription PCR with a viral
RNA-dependent DNA polymerase was performed using the ThermoScript RT-PCR System for
First-Strand Synthesis Kit. According to manufacturer’s specifications, cDNA was synthesized
from 3 g RNA (4.2.1.5.1) with random hexamer primers at 50°C for 50 min in a PTC-225
Thermo Cycler (MJ Research).
4.2.1.5.4 Real-Time PCR reaction
TaqMan was used as a Real-Time PCR system to quantify the transcriptional mRNA level of
genes of interest. The quantification is based on the fluorescence of a reporter dye, whose signal
increases in direct proportion to the amount of amplified PCR product in a sample, that was
detected and quantified.
Quantitative Real-Time PCR was carried out using gene expression assays (4.1.6) or self-
designed primer/probe sets (Table 5) with 900 nM each primer and 250 nM probe. 10 ng cDNA
(4.2.1.5.3) per reaction was used with the FAST qPCR MasterMix Plus following
manufacturer’s instructions scaling the reaction mix down to a total volume of 10 l. To detect
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36
low abundant titin mRNA with the primer/probe sets MEx1-2, MEx6, N2A, and ZEx3, TaqMan
was carried out with the RNA UltraSense One-Step Quantitative RT-PCR System. The provided
manual was adapted to a 10 l reaction including the ROX Reference Dye and 20 ng RNA
(4.2.1.5.1). Amplification reaction for each sample was run in triplicates for 40 cycles using a
7900HT Fast Real-Time PCR System (Applied Biosystems) operating in a 384-well format with
10 l reaction volume.
Data were collected with the Sequence Detection System 2.3 software (Applied Biosystems) and
analyzed using the Comparative CT Method (CT Method) as described in the User Bulletin 2
of the ABI PRISM 7700 Sequence Detection System. For this relative quantification, data were
normalized to 18S cDNA and displayed as fold change relative to the control.
4.2.1.6 Generation of the targeting vector DsRedKI
4.2.1.6.1 Cloning strategy
The targeting vector DsRedKI was designed with the computer program Clone Manager 6 (Sci
Ed Central) spanning the 5′ region of the mouse titin gene around the exon 28. Backbone of the
targeting vector was the vector pGemFRTloxNeo containing the neomycin resistance cassette, in
which the PCR fragments short arm, DsRed, middle arm and long arm (4.2.1.3.1) were
consecutively cloned.
To construct the targeting vector, first the middle arm was cut out from the pGEM-T Easy vector
with the help of the restriction endonucleases Bsp1407I and HindIII. Then the middle arm was
ligated into the vector pGemFRTloxNeo, which was previously linearized with the same
enzymes, leading to MApGemFRTloxNeo. The DsRed fragment subcloned into the pGEM-T
Easy was cut out with Acc65I and cloned into the plasmid MApGemFRTloxNeo opened with
Bsp1407I, to obtain the plasmid DsRedMApGemFRTloxNeo. The short arm of the targeting
vector was gained from the pGEM-T Easy vector by digestion with BamHI and SmaI. The same
enzymes were utilized for digestion of the plasmid DsRedMApGemFRTloxNeo to ligate the
short arm inside resulting in the plasmid DsRedMApGemFRTloxNeoSA. After cutting the long
arm from the pGEM-T Easy vector by XhoI, the final targeting vector DsRedKI was assembled
by ligation of the long arm into the plasmid DsRedMApGemFRTloxNeoSA that was linearized
by XhoI. Before electroporating into ES cells (4.2.3.1), the targeting vector was linearized with
the enzyme ScaI.
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37
A control vector was also cloned as a positive test for the ES cell genotyping PCR (4.2.1.3.3).
Therefore, the DNA fragment short arm control was cut out from the pGEM-T Easy vector by
BamHI and SmaI and ligated into the pGemFRTloxNeo that had been digested with the same
enzymes.
4.2.1.6.2 Enzymatic modifications of DNA
4.2.1.6.2.1 A-tailing
The long arm of the targeting vector was amplified with the Phusion High-Fidelity DNA
Polymerase leading to blunt-ended fragments. To ligate this PCR fragment into the pGEM-T
Easy vector, it was modified using the A-tailing procedure as outlined in the Technical Manual
of the pGEM-T and pGEM-T Easy vector systems. 5.7 l purified long arm PCR fragments
(4.2.1.1.3) were processed for 30 min at 70°C and used afterwards directly for ligation
(4.2.1.6.2.4).
4.2.1.6.2.2 Restriction digest of DNA
For vector linearization and excision of specific DNA fragments, single and double restriction
digests were performed, whereas the amount of restriction endonuclease and buffer as well as
the reaction conditions were set up according to manufacturer’s recommendations. After 2 h of
digestion at 37°C in a volume of 20 l, the linearized vectors were dephosphorylated
(4.2.1.6.2.3) and the DNA fragments were isolated by agarose gel electrophoresis (4.2.1.4).
4.2.1.6.2.3 Dephosphorylation of DNA fragments
To minimize the re-ligation of a vector after digestion with a single enzyme (4.2.1.6.2.2), the
vector was dephosphorylated. 3 U calf intestinal alkaline phosphatase was added to the
linearized vector after the restriction digest and incubated for 30 min at 37°C. The reaction was
stopped by accomplishing agarose gel electrophoresis (4.2.1.4).
4.2.1.6.2.4 Ligation of DNA fragments
PCR products, which were purified (4.2.1.1.3) or purified and A-tailed (4.2.1.6.2.1), were
subcloned into the pGEM-T Easy vector following the instructions of the manufacturer. The
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38
molar ratio of 1:3 for vector to insert was chosen and samples were incubated for 90 min at
room temperature.
For cloning of the targeting vector, DNA fragments were ligated into the vector using the DNA
Ligation Kit according to the protocol provided by the manufacturer. Typically, 20 l buffer A
was combined with 5 l of linearized plasmid vector and DNA insert that were mixed in the
molar ratio of 1:4. 5 l buffer B was added to start the ligation reaction, which was performed at
16°C for 2 h. After the incubation time, DNA ligation mixtures were used directly for
transformation (4.2.2.2).
4.2.2 Microbiological methods
4.2.2.1 Generation of competent bacteria
Competent E.coli DH5 were generated according to a modified “TFB-Based Chemical
Transformation Protocol” published by Hanahan et al. (Hanahan et al., 1991). In step 9 of this
protocol, only DMSO was added to the cell suspension and afterwards 100 l aliquots were
generated, which were stored at -80°C.
4.2.2.2 Transformation of bacteria
Transformation was used to incorporate plasmids into competent E. coli for amplification of
plasmids after ligation. A modified protocol of Hanahan et al. (Hanahan et al., 1991) was
utilized. 200 l of competent bacteria (4.2.2.1) were thawed on ice, mixed with 5 l of ligation
reaction (4.2.1.6.2.4), and incubated for 30 min on ice. Transformation was performed for 45 s at
42°C and afterwards cells were shaken for 40 min at 37°C in 1 ml SOC medium (2% bacto
tryptone, 0.5% yeast extract, 10 mM NaCl, 10 mM MgCl2, 10 mM MgSO4, 2.5 mM KCl,
20 mM glucose, pH 7.0). 100 l of this cell suspension was transferred to a LB-A plate (1%
bacto tryptone, 0.5% yeast extract, 171 mM NaCl, 1.5% agar, 50 g/ml ampicillin, pH 7.0) or to
a LB-A plate coated with 20 l of 2.5 mM X-gal (MBI Fermentas) to identify recombinant
clones by blue-white selection, if the plasmid had the pGEM-T Easy vector as a backbone.
Plates were incubated overnight at 37°C.
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4.2.3 Cell biological methods
4.2.3.1 Gene targeting in mouse ES cells
The ES cell targeting was necessary to incorporate the targeting vector DsRedKI into the
genome of mouse ES cells. It was done by the Transgenic Core Facility of the MDC-Berlin
using the cell line 14.1. This service included culture, storage, and electroporation of ES cells as
well as drug selection, isolation, and passaging of ES cell colonies that were resistant against the
antibiotic G418. These clones were grown in 96-well plates for verification of the targeting
vector within the genome (4.2.1.3.3) and cryopreservation. Homologously recombined ES cells
were expanded and used for formation of embryoid bodies (4.2.3.2) and injection into
blastocysts (4.2.5.2).
4.2.3.2 Formation of embryoid bodies
Embryoid bodies (embryo-like aggregates) were formed from undifferentiated ES cells and
differentiated into beating cardiomyocytes to confirm the integration of the DsRedKI targeting
vector into the titin locus after targeting. The method used was previously described by Wobus
et al. (Wobus et al., 1991). Briefly, ES cells (4.2.3.1), which were positive for the insertion of the
targeting vector tested by PCR, were cultivated in a cell culture incubator at 37°C and 5% CO2
in hanging drops that consisted of 400-600 cells in 20 l differentiation medium (6.7 g DMEM
powder (with L-Glutamine and 4500 mg/l D-Glucose, Invitrogen) and 1.2 g NaHCO3 in 540 ml
ddH2O, after sterile filtration addition of 20% fetal calf serum (FCS, Sigma-Aldrich), 1 x non-
essential amino acids (100x, Invitrogen), 100 U/ml penicillin and 100 g/ml streptomycin
(Invitrogen) and 4.2 μl β-mercaptoethanol). The ES cells aggregated to embryoid bodies within
three days and were then cultivated for five days in suspension using differentiation medium.
Subsequently, single embryoid bodies were picked and transferred to a 24-well plate containing
coverslips that were previously coated with gelatin. The embryoid bodies were allowed to attach
and beating cell clusters were used for immunostaining (4.2.3.4).
4.2.3.3 Preparation and cultivation of primary smooth muscle cells
Smooth muscle cells were isolated from murine aorta by dissection and subsequent enzymatic
treatment of the vascular tissue according to a modified protocol of Ray et al. (Ray et al., 2001).
90 d old control and knockout SMcTiMEx1-2 females were sacrificed by cervical dislocation to
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prepare the aorta. The aorta was transferred to medium (DMEM (4.5 g/l Glucose with
UltraGlutamine I, Lonza) supplemented with 100 U/ml penicillin and 100 g/ml streptomycin
(Invitrogen) and 10% FCS (Sigma-Aldrich)). Using the stereomicroscope Leica MZ7.5, the
adherent fat as well as the tunica adventitia was removed to uncover the tunica media containing
the smooth muscle cells. The tunica media was placed into an enzymatic solution consisting of
1.36 mg collagenase type II and 1 mg eleastase type IV dissolved in 1 ml prewarmed medium
and digested for 1.5 h at 37°C and 5% CO2 in a cell culture incubator to release the smooth
muscle cells. After centrifugation for 3 min at 800 rpm, the enzymatic solution was removed and
each cell pellet was resuspended in 500 l medium and transferred to a 48-well plate coated with
gelatin. In order to allow the cells to attach to wells, the plate was placed into the cell culture
incubator and left undisturbed for five days. For re-plating, cells were washed with PBS
(Cambrex) and treated for 3 min with 0.05% trypsin-EDTA (Invitrogen). After addition of
medium and centrifugation for 3 min at 800 rpm, the cell pellet was resuspended in medium and
seeded. Until usage for immunostaining (4.2.3.4) or the migration assay (4.2.3.5), smooth
muscle cells were allowed to grow to confluence and further passaged.
4.2.3.4 Immunostaining of embryoid bodies and smooth muscle cells
Cell culture medium was removed and embryoid bodies (4.2.3.2) and smooth muscle cells
(4.2.3.3) grown on coverslips were washed once with PBS-azide (0.02% NaN3 in PBS) and
fixed with ice-cold methanol for 15 min. After washing the cells 3 x for 5 min with PBS, they
were permeabilized with 0.2% Triton X-100 in PBS-azide for 15 min and incubated with
blocking solution for 50 min (1% goat serum, 2% BSA in PBS-azide) to stain embryoid bodies
with DsRed as well as -actinin and smooth muscle cells with smooth muscle actin (Table 6) in
PBS-azide overnight at 4°C. Afterwards the cells were washed 2 x 15 min with PBS and
incubated with the secondary antibody (Table 7) diluted in PBS-azide for 1 h. Anti-rabbit Cy3 as
well as anti-mouse Alexa Fluor 488 for the embryoid bodies and anti-rabbit Cy3 for the smooth
muscle cells was used. Subsequently, cells were washed 2 x 15 min with PBS. Nucleus staining
of smooth muscle cells was performed for 30 min with DAPI (1 mg/ml) that was 1:2000 diluted
in PBS. Before the cells, which have attached to coverslips, were placed in Fluorescence
Mounting Medium (Dako) on glass slides, they were washed 2 x for 15 min with PBS and rinsed
3 x with ddH2O. Images were taken using the laser scanning confocal microscope Leica TSP
SP2 and analyzed using Adobe Photoshop CS2.
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4.2.3.5 Cell migration assay
Migration assays were performed using a modified Boyden chamber as described earlier
(Natarajan et al., 1996). This is a filter assay with a cell culture insert that forms two
compartments, which are separated by a porous membrane. Cells can be induced to migrate
from the upper chamber through the membrane into the lower compartment following the
gradient of a chemoattractant.
Primary smooth muscle cells (4.2.3.3) were starved for 24 h in normal medium (DMEM (4.5 g/l
Glucose with UltraGlutamine I, Lonza) supplemented with 100 U/ml penicillin and 100 g/ml
streptomycin (Invitrogen), 0.2% BSA and 0.4% FCS (Sigma-Aldrich)), harvested, washed with
PBS, and resuspended in serum-free medium (normal medium without FCS) to a final
concentration of 106 cells/ml. 400 l of cell suspension was plated on coverslips to document the
presence of smooth muscle cells by immunostaining (4.2.3.4). Additionally, 200 l of each cell
suspension was added to the upper compartment of a cell culture insert containing a
polycarbonate filter with 8 m pores (Corning Incorporated) that was coated with 20 g/ml type
1 collagen (Invitrogen) and placed in a 24-well. The lower compartment of each 24-well was
filled with 600 l serum-free medium supplemented with no or 10-10 M PDGF-B (Reddy et al.,
2003). After 4 h, cells that migrated to the lower side of the filter were fluorescently labeled by
incubation in 450 l serum-free medium with 8 M calcein-AM for 45 min at 37°C and 5%
CO2. Afterwards the cell culture inserts were transferred into 500 l prewarmed 0.05% trypsin-
EDTA (Invitrogen) and incubated for 10 min to detach the migratory cells from the underside of
the membrane. 200 l trypsin-EDTA solution was transferred in duplicates into a flat-bottom
black 96-well plate (Greiner Bio-One GmbH) to quantify the cell count of each sample by
reading the fluorescence at an excitation of 485 nm and an emission of 520 nm in a Synergy HT
Multi-Mode Microplate Reader (BioTek).
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4.2.4 Biochemical methods
4.2.4.1 Protein preparation
4.2.4.1.1 Protein lysates for 2D-gel electrophoresis
Uterine tissue of E3.5 pregnant SMcTiMEx1-2 mice (4.2.5.7) was ground under liquid nitrogen
and mixed with 4 x volume of 2D-lysis buffer (8 M urea, 2 M thiourea, 4% chaps, 0.5%
pharmalyte, 10 mM DTT, 1 mM PMSF, 100 nM Calyculin A). After sonication using the
sonopuls sonicator SH70G (Brandelin electronic GmbH) with 70% power for 2 min on ice, the
cell debris was spun down at 50,000 rpm at 4°C for 30 min. The protein concentration of the
supernatant was determined by the Bradford assay (4.2.4.2.1) and the protein lysates were stored
in aliquots at -80°C.
4.2.4.1.2 Protein lysates for VAGE
Protein lysates of heart, quadriceps, and soleus of TiEx28DsRed mice in estrous stage (4.2.5.7)
were prepared according to a modified protocol of Warren et al. (Warren et al., 2003). Tissues
were pulverized under liquid nitrogen and transferred with the help of 40 x volume titin sample
buffer (8 M urea, 2 M thiourea, 3% SDS, 0.05 M Tris-HCl, 0.03% bromophenol blue, 75 mM
DTT, pH 6.8) into a Dounce homogenizer (Wheaton). After 3 min of homogenization, samples
were vortexed thoroughly, left at room temperature for 25 min, and centrifuged for 5 min at
13,000 rpm. Supernatants were removed and their protein concentration was measured using the
amido black method (4.2.4.2.2). Afterwards, supernatants were aliquoted and stored at -80°C.
4.2.4.1.3 Protein lysates for multiplex bead immunoassay
Tissue Extraction Reagent I (Invitrogen) was supplemented with protease inhibitor cocktail
(Sigma-Aldrich) according to manufacturer’s manual. Uteri of E3.5 pregnant SMcTiMEx1-2
mice (4.2.5.7) were homogenized in 1 ml buffer per 100 mg tissue with the help of an Ultra-
Turrax T-8 (IKA-Werk). Afterwards, samples were centrifuged for 5 min at 10,000 rpm and 4°C
to pellet the tissue debris. Before aliquoting and storing at -80°C, the protein concentration of
the supernatant was analyzed using the Bradford method (4.2.4.2.1).
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4.2.4.1.4 Protein lysates for ELISA
Uterine horns of SMcTiMEx1-2 mice pregnant at E3.5 (4.2.5.7) were homogenized in 0.5 ml of
50 mM phosphate buffer (0.16% KH2PO4, 1.04% Na2HPO4 x 7 H2O in ddH2O, pH 7.4) using an
Ultra-Turrax T-8 (IKA-Werk). After centrifugation of the homogenates at 10,000 g for 20 min at
4°C, the supernatant was stored in aliquots at -80°C until assayed directly using ELISA (Li et
al., 2004).
4.2.4.2 Protein quantification
4.2.4.2.1 Bradford assay
Protein amount was quantified with the method of Bradford (Bradford, 1976), which is based on
the absorbance maximum shift of Coomassie Brilliant Blue G-250 from 465 nm to 595 nm in
response to protein binding. The Bradford protein assay (Bio-Rad) was carried out according to
manufacturer’s instruction using the Ultrospec 2100 pro UV/Visible Spectrophotometer (GE
Healthcare).
4.2.4.2.2 Amido black method
Protein concentration of VAGE samples was determined with the amido black method because
SDS in the lysis buffer impairs the Bradford method (Schaffner and Weissmann, 1973). This
quantification is predicated on the binding of the dye amido black to protein that is bound on a
membrane. BSA (bovine serum albumin) dilutions in VAGE lysis buffer, which were used to
generate a standard curve, and 1 l of each protein sample were spotted on a nitrocellulose
membrane (GE Healthcare) in duplicates. On an orbital shaker, the membrane was incubated for
1 min in amido black staining solution (45% methanol, 10% acetic acid, 0.1% amido black)
followed by destaining (90% methanol, 2% acetic acid) until the background staining of the
membrane was removed. The membrane was cut to separate the protein spots and to transfer
each protein spot to an individual reaction tube containing 800 l elution buffer (50% ethanol,
0.05 mM Na2EDTA, 25 mM NaOH). During 30 min incubation on a shaker, the protein spots
were discolored releasing the bound amido black into the elution buffer. The absorbance of
amido black in the elution buffer was measured at 630 nm using the Ultrospec 2100 pro
UV/Visible Spectrophotometer (GE Healthcare). The protein concentration in each sample was
determined with the help of the BSA standard curve.
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4.2.4.3 Protein gel electrophoresis
4.2.4.3.1 2D-gel electrophoresis
4.2.4.3.1.1 First dimension
In the first dimension, proteins of a lysate were separated according to their isoelectric point.
18 cm immobilized pH gradient (IPG) strips with a pH-gradient of 3 to 10 (GE Healthcare) were
rehydrated overnight at room temperature in 340 l rehydration solution (8 M urea, 2 M
thiourea, 2% chaps, 0.5% Pharmalyte pH 3-10 (GE Healthcare), 12 l/1ml DeStreak-reagent
(GE Healthcare), 0.03% bromophenol blue). 125 g protein lysate (4.2.4.1.1) were replenished
with rehydration solution to a total volume of 50 l and applied to IPG strips by cup-loading.
The samples were isoelectric focused under DryStip Cover Fluid (GE Healthcare) at 20°C using
the MultiphorII (GE Healthcare) with the following profile:
1. gradient 300 V 30 min
2. step 300 V 30 min
3. gradient 500 V 1 h
4. step 500 V 1 h
5. gradient 3500 V 1.5 h
6. step 3500 V 5.5 h
After focusing, the strips were immediately used for the second dimension.
4.2.4.3.1.2 Second dimension
After isoelectric focusing, the second dimension was used to separate proteins dependent on
their molecular weight using continuous sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (SDS-PAGE). Each IPG strip was incubated for 15 min at room temperature in
10 ml equilibration buffer A (6 M urea, 50 mM Tris-HCl, 30% glycerol, 2% SDS, 22 mM DTT)
and afterwards for 15 min at room temperature in 10 ml equilibration buffer B (6 M urea,
50 mM Tris-HCl, 30% glycerol, 2% SDS, 0.135 M iodoacetamide, 0.03% bromophenol blue) on
an orbital shaker. Then the IPG strips were placed on SDS-gels (T=12.5%, C=2.6%; 133 ml
ddH2O, 152 ml 40% acrylamide, 85 ml 2% bisacrylamide, 125 ml 1.5 M Tris-base pH 8.8, 5 ml
10% SDS, 1.25 ml 10% ammonium persulfate, 0.25 ml TEMED) and overlaid with 0.5 %
agarose in 1 x SDS running buffer (25 mM Tris-base, 192 mM glycine, 0.1% SDS). For SDS-
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PAGE, 1 x SDS running buffer in the lower chamber and 2 x SDS running buffer in the upper
chamber of an Ettan Dalt Twelve Electrophoresis Unit (GE Healthcare) was used.
Electrophoresis was performed at 16°C with 2 W/gel for 30 min followed by 15 W/gel.
Afterwards, the gels were stained with Coomassie (4.2.4.4).
4.2.4.3.1.3 Quantification
To visualize protein differences in uteri of knockout compared to control mice, gels were
analyzed with the Delta2D Version 3.4 software (DECODON). The percent volume value (% V)
of each protein spot for six knockout and six control samples was averaged and the quotient of
average % V knockout and average % V control was calculated for each spot. The resulting ratio
values were filtered to obtain the proteins that were downregulated (ratio < 0.8) and upregulated
(ratio > 1.3) in knockout compared to control uteri. Weak protein spots with an average % V of
lower than 0.03 in knockout and control gels were excluded.
4.2.4.3.1.4 Mass spectrometry
To identify proteins by mass spectrometry according to a published procedure (Raddatz et al.,
2008), protein spots of 2D-gels stained with Coomassie Brilliant Blue were cut out and send to
the Ernst-Moritz-Arndt-University of Greifswald (group of Professor Dr. Hecker).
4.2.4.3.2 Vertical SDS-agarose gel electrophoresis (VAGE)
Proteins with a high molecular weight can only be poorly separated using SDS-PAGE. Hence a
vertical SDS-agarose gel electrophoresis (VAGE) with a 1% agarose gel was performed to
separate titin isoforms of heart, quadriceps, and soleus protein lysates (4.2.4.1.2). VAGE was
adapted from a published protocol (Warren et al., 2003). A SE 600 Vertical Gel Unit (GE
Healthcare) with VAGE running buffer (50 mM Tris-base, 0.384 M glycine, 1% SDS) in the
lower chamber and VAGE running buffer supplemented with 10 mM -mercaptoethanol in the
upper chamber was used. SDS-agarose gels (1% SeaKem Gold Agarose (Cambrex), 30%
glycerol, 50 mM Tris-base, 0.384 M glycine, 1% SDS) were run at 15 mA constant current in a
4°C cold room. After electrophoretic separation, gels were stained with Coomassie Brilliant
Blue (4.2.4.4) or transferred to a membrane (4.2.4.5.1).
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4.2.4.4 Coomassie staining
Each step of the staining procedure was conducted on an orbital shaker. 2D-gels were fixed
(50% ethanol, 10% acetic acid) for 1 h and gels from VAGE were first fixed (50% methanol,
12% acetic acid, 5% glycerol) for 1 h and then dried at 37°C overnight to minimize background
staining. Afterwards both gel types were washed 3 x for 10 min with ddH2O followed by
overnight staining with a Coomassie Brilliant Blue G-250 (Sigma-Aldrich) solution prepared
according to Neuhoff et al. (Neuhoff et al., 1988). Excess dye was removed by ddH2O washing
and gel images were taken using the Camilla or Stella 8300 imaging system (Raytest). VAGE
gels were processed using the Aida Image Analyzer v. 4.24 software (Raytest) and 2D-
electrophoresis gels were analyzed using the Delta2D software (4.2.4.3.1.3).
4.2.4.5 Western blotting
4.2.4.5.1 Semi-dry blotting
The semi-dry electroblotting was used to transfer titin for immunological detection to a PVDF
(polyvinylidene difluoride) membrane (GE Healthcare). The membrane was incubated 10 s in
100% methanol, 2 min in ddH2O and 15 min in anode buffer II (25 mM Tris-base, 0.05% SDS,
10% methanol, pH 10.4). VAGE-gels (4.2.4.3.2) were equilibrated for 10 min in cathode buffer
(25 mM Tris-base, 0.05% SDS, 10% methanol, 40 mM caproic acid, 10 mM -mercaptoethanol,
pH 10.4) and then assembled with the membrane and Whatman 3MM Chr paper (Schleicher &
Schuell). Previously, the blotting paper towards the cathode was wet in cathode buffer and the
blotting paper towards the anode was soaked in anode buffer I (300 mM Tris-base, 0.05% SDS,
10% methanol, pH 10.4) and anode buffer II. The transfer was performed at constant 250 mA for
90 min in the PerfectBlue Semi-Dry Electroblotter SEDEC M (PEQLAB).
The blot transfer efficiency was controlled by incubation for 3 min with Ponceau S staining
solution (Sigma-Aldrich). Excessive dye was removed with ddH2O to make the protein bands
visible. Then the membrane was destained with PBS-T (0.1% Tween 20 in PBS).
4.2.4.5.2 Immunological detection of proteins
The detection of specific proteins after semi-dry Western blotting was done by an
immunological reaction using antibodies. Unspecific binding sites on the membrane were
blocked with 5% milk powder in PBS-T for 1 h followed by incubation of the membrane with
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the primary antibody (Table 6) in 5 % protease free BSA (SERVA) in PBS at 4°C overnight.
After washing the membrane 3 x 15 min with PBS-T, it was incubated with the secondary
antibody (Table 7) in 5% milk powder in PBS for 1 h and washed 2 x 15 min with PBS-T and
1 x 15 min with PBS. The secondary antibody was conjugated with horseradish peroxidase and
imaged with the ECL-system (enhanced chemiluminescence) using the SuperSignal West Femto
Maximum Sensitivity Substrate (Thermo Scientific) according to manufacturer’s instruction.
Chemiluminescence was detected with the Stella 8300 Imaging system (Raytest).
4.2.4.6 Histology
4.2.4.6.1 Immunostaining
Heart and quadriceps of TiEx28DsRed animals in estrous stage (4.2.5.7) were fixed overnight at
4°C in formalin. Then the tissue was dehydrated in 30% sucrose in PBS overnight at 4°C and
embedded in Tissue-Tek O.C.T. Compound (Sakura). 10 m thick longitudinal sections were cut
using the Cryostat HM 560 Cryo-Star (Thermo Scientific) with a specimen temperature of -25°C
and a knife temperature of -35°C and placed on HistoBond slides (Marienfeld). Cryosections
were stored at -20°C.
For immunostaining, sections were air-dried, fixed for 15 min with 2% paraformaldehyde in
PBS-azide (0.02% NaN3 in PBS), and washed 2 x for 5 min with PBS. After permeabilization
and blocking for 1 h (10% goat serum, 0.3% Triton X-100, 0.2% BSA in PBS) samples were
washed 4 x with PBS for 5 min and incubated with the primary antibody titin M8/M9 (Table 6)
in PBS at 4°C in a wet chamber overnight. Excess primary antibodies were removed by a 6 x
PBS wash each for 10 min. Sections were stained with an anti-rabbit IgG biotin-conjugated
antibody (Table 7) in PBS for 1 h followed by 6 x PBS washing. Then the signal of the titin
M8/M9 antibody was amplified by incubation with streptavidin conjugated to Alexa Fluor 647
(1:1000; Invitrogen) in PBS for 1 h. Finally, sections were washed 4 x for 5 min with PBS and
2 x for 1 min with ddH2O and mounted in Fluorescence Mounting Medium (Dako). Protein
localization was documented using the laser scanning confocal microscope Leica TSP SP2 and
images were analyzed using Adobe Photoshop CS2.
4.2.4.6.2 LacZ staining
LacZ staining was performed to monitor the activity of the SMMHC cre-recombinase in uteri of
SMcGtROSA26 reporter mice. These mice contain a lox-flanked DNA stop element followed by
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the lacZ gene knocked into the ROSA26 locus. Only if the floxed stop segment is excised by
cre-recombinase activity, the lacZ is expressed. The ability of the -galactosidase to cleave X-
gal leading to a blue insoluble product is used to monitor the presence of this enzyme. The
staining procedure for uteri of SMcGtROSA26 mice in estrus stage (4.2.5.7) was adapted from
the protocol “Staining Frozen Sections” (Nagy et al., 2003b). After consecutive incubation in
fixative (0.2% paraformaldehyde, 2 mM MgCl2, 5 mM EGTA, 0.1 M PIPES, pH 6.9) and 30%
sucrose in PBS-M (2 mM MgCl2 in PBS) at 4°C overnight, the tissue was embedded in Tissue-
Tek O.C.T. Compound (Sakura) to cut cross sections as described before (4.2.4.6.1).
The samples were air-dried and fixed for 5 min with 2% paraformaldehyde and 0.125%
glutaraldehyde in PBS followed by a 3 x PBS-M wash for 1 min. Afterwards the slides were
incubated 3 x for 2 min in detergent solution (2 mM MgCl2, 0.01% Na-deoxycholate, 0.02%
NP40 in PBS) and 2 min in staining solution (2 mM MgCl2, 5 mM potassium ferricyanide,
5 mM potassium ferrocyanide in PBS). Sections were stained overnight at 37°C with 2.5 mM X-
gal (MBI Fermantas) in staining solution. For the counterstain with eosin, the samples were
rinsed in ddH2O for 2 min and stained for 1 min with eosin solution (1% eosin, 0.05% acetic
acid in ddH2O). Before embedding in Roti-Histokitt (Roth), dehydration of the sections was
performed with increasing alcohol concentrations (70%, 90%, and 100%) each for 30 s. Pictures
were taken using the BX51 Research Microscope (Olympus).
4.2.4.7 ELISA
An enzyme-linked immunosorbent assay (ELISA) kit was used to measure the metabolite 12-
hydroxyeicosatetraenoic acid (12-HETE) in homogenates of uterine samples (4.2.4.1.4)
according to the manufacturer’s instructions. The principle of this method is based on the
competitive binding of 12-HETE, which is conjugated to alkaline phosphatase, and the
metabolite in the sample to an antibody that is immobilized on wells. The obtained signal read at
405 nm in a Synergy HT Multi-Mode Microplate Reader (BioTek) was indirectly proportional to
the amount of 12-HETE in the sample. Data acquired for the standards were displayed as logit-
log plot and a linear regression fit was performed to obtain a standard curve, which was used to
calculate the concentrations of 12-HETE in the sample. Metabolite concentration was
normalized to the total protein concentration of the sample that was measured using the
Bradford method (4.2.4.2.1).
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4.2.4.8 Multiplex bead immunoassay
Multiplex bead immunoassay is an immunoassay on the surface of fluorescent-coded beads,
which allows the investigation of several analytes in a single sample at the same time. With the
help of this method, the growth factors G-CSF (granulocyte colony-stimulating factor), FGF
(fibroblast growth factor) basic, PDGF-B, and VEGF (Growth Factor Mouse 4-Plex Panel) as
well as IL-1β (interleukin-1β), IL-6, IL-15, IP-10 (interferon-inducible protein 10), LIF, MCP-1
(monocyte chemotactic protein-1), and TNF-α (tumor necrosis factor-alpha; Mouse
Cytokine/Chemokine Panel-7-Plex) were quantitatively determined in serum (4.2.5.6) and
uterine tissue lysate of E3.5 pregnant mice (4.2.4.1.3). Prior to immunoassay analysis, all tissue
lysates were normalized against their total protein concentration. The kits were carried out
following the protocol given by the companies utilizing the Luminex 200TM instrument with the
software xPONENT 3.0 (Luminex Corporation). The concentrations of the analytes in the sera
or lysates were calculated from the standard curve using as curve fitting software
xPONENT 3.0. Thereby the four parameter algorithm for the Invitrogen kit and the 5-parameter
logistic for the Millipore kit was used.
4.2.5 Animal procedures
4.2.5.1 Maintenance of the mouse colony
TiEx28DsRed, FLP-transgene, SMMHCcre, SMcGtROSA26, TiMEx1-2, and SMcTiMEx1-2
mice were derived from own breeding. The founder of the transgenic mouse lines FLP-
recombinase (FLP-transgene) and smooth muscle myosin heavy chain (SMMHC) cre-
recombinase (SMMHCcre) were kindly provided (Rodriguez et al., 2000; Xin et al., 2002).
SMMHCcre males were bred with 129S-Gt(ROSA)26Sortm1Sor/J reporter mice, which were
purchased from The Jackson Laboratory, to obtain the double heterozygous strain
SMcGtROSA26. The conditional knockout strain of titin’s M-band exons 1 and 2 (TiMEx1-2)
was available in the group (Gotthardt et al., 2003), so that the smooth muscle specific TiMEx1-2
(SMcTiMEx1-2) strain was obtained by breeding the SMMHCcre strain with TiMEx1-2 mice.
The TiEx28DsRed mouse strain was newly generated (4.2.5.3). All mice were numbered by ear
marks and tail biopsies were taken for genotyping (4.2.1.3.3 and 4.2.1.3.4).
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All mice used in the studies were of 129/S6 (Taconic) and C57Bl/6 (Charles River) mixed
background. They were housed in the animal facility of the MDC-Berlin in a 12 h light/dark,
20°C temperature controlled environment with free access to food and water using individually
ventilated cages (IVCs). The animal procedures were performed in accordance to the regulatory
rules for Animal Welfare of the German Society for Laboratory Animal Science.
4.2.5.2 Generation of chimeric mice
The generation of mouse chimera was done by the Transgenic Core Facility of the MDC-Berlin.
This process included the microinjection of ES cells from one clone (4.2.3.1), which
incorporated the targeting vector DsRedKI within the titin locus, into blastocysts obtained from
superovulated C57Bl/6 mice. These blastocysts were surgically transferred into pseudopregnant
recipient mice leading to chimeric offspring. After weaning, the chimeras were transferred to the
animal colony for breeding (4.2.5.3).
4.2.5.3 Chimera breeding and generation of the TiEx28DsRed strain
Chimeras (4.2.5.2) were mated with C57Bl/6 (Charles River) mice to obtain offspring that
carries the DsRedKI targeted gene. Offspring with brown coat color (agouti) that contained the
targeted allele was bred with FLP-transgene mice (Rodriguez et al., 2000) to delete the
neomycin resistance cassette. Afterwards the FLP-transgene was removed from double
heterozygous mice by crossing them with 129/S6 (Taconic) animals. Mice heterozygous for the
altered titin locus were used to generate the colony of the TiEx28DsRed strain.
4.2.5.4 Staging of the estrous cycle
Induced by ovarian hormones, the murine reproductive system undergoes periodic physiological
changes, which are classified into the four stages metestrus, diestrus, proestrus and estrus (Allen,
1922; Snell, 1941). Each stage shows a distinct uterine and vaginal cytology. To collect non-
pregnant uteri in the estrous stage of the estrous cycle and to determine the estrous cycle length,
it was necessary to monitor changes in vaginal epithelial cells at 24 h intervals by vaginal lavage
(Frick and Berger-Sweeney, 2001). Therefore, the vagina of 90 d old wild-type, heterozygous,
and homozygous TiEx28DsRed, double heterozygous and heterozygous SMcGtROSA26 mice,
SMMHCcre mice, and control, heterozygous, and knockout SMcTiMEx1-2 female mice was
rinsed with ddH2O to extract vaginal cells and the fluid was placed onto a microscope slide.
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After evaporation of the water, the sample was examined under the inverted light microscope
Olympus CK40 applying the guidelines of Allen and Snell (Allen, 1922; Snell, 1941). At least,
two continuous cycles were observed before sacrificing the mice.
4.2.5.5 Timed mating
90 d old control, heterozygous and knockout females of the SMcTiMEx1-2 strain as well as
SMMHCcre mice were mated. Thereby knockout males were used for control females and
control males were used for heterozygous and knockout females to get embryos with similar
genotypes. SMMHCcre females were bred with 129/S6 males. Mating was set up at 6 pm and
the morning of detecting a vaginal plug was designated as E0.5.
4.2.5.6 Collection of serum
The retrobulbar venous plexus puncture was used to obtain blood samples from E3.5 pregnant
control and knockout SMcTiMEx1-2 mice (4.2.5.5). Mice were anesthetized with ether and
blood was withdrawn using a capillary tube that was directed towards the major venous structure
of the eye. For serum collection, blood was allow to coagulate for 5 min at room temperature
and then 55 min on ice before it was centrifuged for 15 min at 3000 rpm. The upper serum phase
was taken, frozen in liquid nitrogen and stored at -80°C until investigation by multiplex bead
immunoassay (4.2.4.8).
4.2.5.7 Tissue harvesting
The body weight of mice in estrous stage (4.2.5.4) and of E3.5 pregnant control and knockout
SMcTiMEx1-2 mice (4.2.5.5) was measured. Animals were killed by cervical dislocation to
dissect uterus from SMcGtROSA26, SMMHCcre as well as SMcTiMEx1-2 mice and heart,
quadriceps, and soleus from TiEx28DsRed mice. The tissue weight of uterus and heart was
determined and if applicable the left uterine horn was flushed with DEPC-PBS to confirm the
pregnancy day E3.5 by the presence of blastocysts. Then, the tissue was used for cryosections
(4.2.4.6) or quick frozen in liquid nitrogen and stored at -80°C until RNA or protein preparation.
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4.2.5.8 Counting of blastocysts
Control and knockout SMcTiMEx1-2 mice pregnant at E3.5 (4.2.5.5) were killed by cervical
dislocation to take out the uteri. Blastocysts were flushed from both uterine horns with PBS and
counted using the stereomicroscope Leica MZ7.5 (Ye et al., 2005).
4.2.5.9 Evans Blue staining
To determine the number and localization of embryo implantation sites at E4.5 and E5.5, they
were stained by Evans Blue. This is an azo compound dye that binds to serum albumin in the
blood thereby making areas of vascular permeability, such as implantation sites, visible. Timed
mated SMMHCcre as well as control, heterozygous and knockout SMcTiMEx1-2 females
(4.2.5.5) were intravenously injected with 150 l Evans Blue (1% in PBS) solution (Ye et al.,
2005). 5 min after injection, mice were killed by cervical dislocation and uterus and heart were
dissected. Their weight was determined and pictures of the uteri were taken using an Olympus
720 SW camera. Afterwards uteri horns of E4.5 pregnant females were investigated for the
presence of blastocysts as described (4.2.5.8).
4.2.5.10 Angiogenesis study
A standard method to investigate angiogenesis in vivo is the injection of matrigel. This is a
solubilized basement membrane preparation extracted from Engelbreth-Holm-Swarm mouse
sarcoma that is a tumor rich in extracellular matrix proteins. The matrigel plug assay was carried
out as described previously with modifications (Isaji et al., 1997). Matrigel Phenol Red-free (BD
Biosciences) was thawed overnight at 4°C and mixed with 30ng/ml VEGF and 64 units/ml
heparin. 0.5 ml of this mixture was injected subcutaneously in one site at the abdominal midline
of 90 d old male and female control and knockout SMcTiMEx1-2 mice. Upon injection, the
matrigel hardened to form a plug. After seven days, mice were euthanized by ether and the
matrigel was harvested, weighted, and frozen in liquid nitrogen. Afterwards 150 l blood was
taken by cardiac puncture, mixed with 7.53 l heparin (64 units/ml) and frozen in liquid
nitrogen. Matrigel as well as the blood were stored at -80°C until analyzing for the formation of
blood vessels by determination of hemoglobin content. Therefore, matrigel plugs were digested
with 2 mg/ml collagenase type I and 2 mg/ml dispase II in TESCA buffer (50 mM TES,
0.36 mM CaCl2, pH 7.4 at 37°C) for 2 h at 37°C and 750 rpm and afterwards the quantity of
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hemoglobin was measured using Drabkin’s reagent (Sigma-Aldrich) according to
manufacturer’s instructions. Blood hemoglobin content was also determined with Drabkin’s
reagent to express the results as blood volume per unit weight of matrigel (Bandyopadhyay et
al., 2002).
4.2.6 Statistical analysis
Evaluation of data was done using the GraphPad Prism 5 software (GraphPad Software). Data of
one group were displayed as mean with standard error of the mean (SEM). The t-test was
applied to compare the means of two groups. The comparison of the means of three and more
unmatched groups was done with the help of the one-way ANOVA. The two-way ANOVA was
used to determine how experimental data are affected by two factors, both individually and
together. For the 2D-gel analysis, a P-value of P<0.1 * fulfilled the significance criteria and for
the other experiments a P-value of P<0.05 *, P<0.01 **, and P<0.001 *** was considered to be
significant.
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5 Results
To investigate titin in smooth muscle, we used a morphological and a functional approach. A
knockin mouse model was established to visualize smooth muscle titin with the help of the red
fluorescence protein DsRed. Mice expressing the titin-DsRed were viable and fluorescence
emitted by the DsRed was detected in muscle tissue. Furthermore, a loss of function approach
was used to analyze the role of titin’s kinase region, which is encoded by titin’s M-band exons 1
and 2, in smooth muscle. Deletion of titin’s kinase region in uterine tissue led to an embryo
implantation phenotype with impaired angiogensis and tissue remodeling.
5.1 The DsRedKI mouse model
5.1.1 Cloning of the targeting vector DsRedKI
We cloned a targeting vector to insert the red fluorescence protein DsRed into titin. The DsRed
should be present close to titin’s N-terminus but not interfere with titin’s integration into the Z-
disc of the sarcomere and with the protein binding sites of T-cap, sANK, -actinin, and obscurin
(Figure 5). Hence the DsRed coding sequence was incorporated into the exon 28 of titin
considering its domain composition. This exon encodes unique sequences and Ig-domains (Bang
et al., 2001a), which are located at titin’s Z-disc to I-band junction (Figure 4). In the targeting
vector DsRedKI, the coding sequence of the DsRed was inserted between the coding sequence
of the Ig-domain Z9 and the Ig-domain I1 of titin’s Z/I-band junction preserving the reading
frame (Figure 7). The whole targeting vector spanned the genomic region from titin’s exon 18 to
exon 31. It consisted of the long arm from exon 18 to exon 28 until the DsRed insertion, the
DsRed, the middle arm, the neomycin resistance cassette flanked by two FRT-sites, and the short
arm. The middle arm included remains of exon 28 and intron sequence and the short arm
contained intron sequence as well as exon 29 to exon 31. The targeting strategy included
insertion of the targeting vector, which was linearized with the enzyme ScaI, into ES cells using
electroporation. The integration of the targeting vector into the titin wild-type allele of ES cells
by homologous recombination led to the targeted allele. After germline transmission, the
neomycin resistance cassette was removed by the enzyme FLP-recombinase resulting in mice
carrying the knockin allele (recF). Compared to the wild-type allele, the KI allele contained one
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FRT-site in an intron and the coding sequencing of the DsRed in exon 28 within the reading
frame.
Figure 7: Targeting strategy to insert the red fluorescence protein DsRed. The illustration shows the
exon/intron (boxes/lines) structure of the titin gene at the Z/I-junction of the sarcomere spanning the region from
exon 18 to exon 31. In the targeting vector, the DsRed coding sequence was inserted into the exon 28. Homologous
recombination of the targeting vector, which was linearized with the enzyme ScaI, into the wild-type (WT) allele in
ES cells led to the targeted allele (neo). After germline transmission, the neomycin resistance cassette (neo) flanked
by two FRT-sites (FRT) was removed by the FLP-recombinase leading to mice carrying the knockin allele (recF)
including the DsRed and one FRT-site. Primers for genotyping are indicated as arrows. The coding regions for the
Ig domains (red) and the unique sequences (blue) are shown for the exon 28 and the Ig domains Z9 and I1 are
specified.
The targeting vector DsRedKI was assembled using a PCR-based strategy, in which the PCR
fragments were cloned into a vector containing the neomycin resistance cassette (supplement
Figure 30). Verification of the targeting vector was done by restriction digest with BamHI,
Bsp1407I, NcoI, and ScaI leading to DNA fragments in the predicted sizes (Figure 8; BamHI
8108 bp, 4618 bp, 2621 bp; Bsp1407I 10346 bp, 4378 bp, 623 bp; NcoI 11301 bp, 3223 bp,
823 bp; ScaI 15347 bp).
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Figure 8: Verification of the targeting vector DsRedKI. Restriction digest with BamHI, Bsp1407I, NcoI, and
ScaI show the predicted fragment sizes (BamHI 8108 bp, 4618 bp, 2621 bp; Bsp1407I 10346 bp, 4378 bp, 623 bp;
NcoI 11301 bp, 3223 bp, 823 bp; ScaI 15347 bp) on a DNA-agarose gel. Uncut targeting vector was used as
control.
The coding regions and the neomycin resistance cassette including the FRT-sites were sequenced
to ensure that no mutations have occurred. The unique ScaI restriction site within targeting
vector and was used for linearization before the ES cell targeting.
5.1.2 Generation of the titin-DsRed knockin mouse strain TiEx28DsRed
Two gene targeting experiments were done to insert the targeting vector by homologous
recombination into the genome of mouse ES cells resulting in 384 clones that were screened by
PCR for the integration of the targeting vector into the titin locus. This ES cell genotyping PCR
was established with the help of the cloned control vector, which was used as template in
different amounts (Figure 9A). The sensitivity of the ES cell genotyping PCR reached 100 fg
with plasmid DNA, but a very weak signal was also noticeable at 15 fg.
Figure 9: ES cell genotyping by PCR. A) Establishing the PCR using the control vector in different amounts as
indicated. This PCR is sensitive to detect 100 fg of plasmid DNA as visible by the 1971 bp PCR product of the
expected size on an agarose-gel. A very weak PCR-signal was also noticeable at 15 fg. B) The ES cell clones E3
and F5 were identified out of 384 clones by PCR for having incorporated the targeting vector into the titin gene by
homologous recombination. Thereby, three different concentrations of DNA as indicated as well as the control
vector as positive control (+) were used for the PCR.
The ES cell genotyping PCR indentified the targeted ES cell clones E3 and F5 for having
incorporated the targeting vector into the titin gene (Figure 9B). To test the expression of the
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DsRed within the titin gene and the proper incorporation of the titin-DsRed into the sarcomere,
ES cells from the clones E3 and F5 were differentiated into cardiomyocytes via embryoid
bodies. 17 days after forming aggregates, cell clusters beat indicating the presence of
cardiomyocytes in the culture. Since it was not possible to detect fluorescence emitted by the
DsRed, the cells were stained using an antibody against DsRed. Additionally, a -actinin
antibody was used to confirm the localization of the red fluorescence protein close to the Z-disc
of the sarcomere (Figure 10).
Figure 10: Proper localization of titin-DsRed in the sarcomere. Immunostaining using an antibody against
DsRed (red) and -actinin (green) of embryoid bodies obtained from the ES cell clones F5 and E3 that were
differentiated into beating cardiomyocytes (n=3). For clone F5, DsRed colocalized with -actinin (yellow, overlay)
indicating that DsRed is present at the Z-disc close to titin’s N-terminus, which is incorporated properly into the
sarcomere leading to a striated pattern. The presence of cardiomyocytes obtained from clone E3 was confirmed by
the -actinin staining, but it was not possible to detect the DsRed.
The striated pattern of the DsRed staining (red) showed the proper incorporation of the titin-
DsRed into the sarcomere of cardiomyocytes derived from the ES cell clone F5. The presence of
the DsRed at the Z-disc was demonstrated by its colocalization (yellow, overlay) with the Z-disc
protein -actinin (green). It was not possible to detect DsRed antibody staining in cells of the ES
cell clone E3, although the striated pattern of -actinin confirmed the presence of
cardiomyocytes. Since cardiomyocytes derived from the clone F5 expressed titin-DsRed, ES
cells from this clone were injected into blastocysts to generate chimeric mice. Two blastocyst
injections resulted in nine chimeras. Five chimeric animals had a brown coat color of more than
70% indicating a high contribution from the ES cell clone and a low contribution of the host
C57BL/6 blastocyst, which would lead to a black fur. All chimeras were bred with C57BL/6
mice and thereby germline transmission of the targeted allele was obtained. Genotyping of mice,
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which were used to establish and maintain the TiEx28DsRed strain, was done by PCR (Figure
11) with primers as indicated in Figure 7. Mice that show PCR products in the expected sizes
were used for further breeding.
Figure 11: Genotyping of TiEx28DsRed animals. PCR-analysis of the possible genotypes (wild-type +/+ FLP-;
heterozygous neo/+ FLP- and R/+ FLP+ or FLP-; homozygous R/R FLP-; neo: neomycin resistance cassette; R:
DsRed) revealed, that neo/+ FLP- mice incorporated the targeting vector with neomycin resistance cassette in the
titin gene (neo-PCR 1971 bp) including the coding sequence for the DsRed (DsRed-PCR 693 bp). To remove the
neomycin resistance cassette, these animals were mated with FLP-transgene mice leading to double heterozygous
R/+ FLP+ mice. They contain the FLP-transgene (FLP-PCR 480 bp) as well as a recombined targeted allele (recF
369 bp) and a titin wild-type allele (WT 282 bp). In the next generation, the FLP-transgene was crossed out leading
to R/+ FLP- mice, which were used for breeding to establish the TiEx28DsRed strain colony resulting also in
homozygous knockin (R/R FLP-) and wild-type control mice (+/+ FLP-).
The first sign of germline transmission was the appearance of pups with brown coat color, which
were investigated for the presence of the DsRedKI targeting construct in the titin gene. This was
done with one primer binding within the neomycin resistance cassette and the other binding in
the titin gene outside of the genomic region present in the targeting vector (neo-PCR). These
animals were additionally genotyped for the presence of the DsRed coding sequence (DsRed-
PCR). Since the neomycin resistance cassette can affect the phenotype of genetically engineered
mice (Kaul et al., 2000), heterozygous animals carrying the targeting vector (neo/+ FLP-) were
bred with FLP-transgene mice leading to double heterozygous mice (R/+ FLP+; DsRed and
FLP-PCR). The deletion of the neomycin resistance cassette by the FLP-recombinase was
controlled by the recF-PCR resulting in a recF as well as a wild-type (WT) band. Both PCR
products were formed because recombination had occurred in heterozygous animals that carry
also one wild-type allele. Animals carrying heterozygous the targeting vector (neo/+ FLP-) show
a wild-type signal in the recF-PCR as well. In the next generation, the FLP-transgene was
crossed out to obtain heterozygous animals (R/+ FLP-; DsRed and recF-PCR). They were used
to generate the colony of the TiEx28DsRed strain. Thereby, also homozygous knockin animals
(R/R FLP-) that show only the recF PCR product as well as wild-type controls (+/+, FLP-) that
show only the wild-type PCR product were born (DsRed and recF-PCR). Animals obtained by
heterozygous breeding were used for analysis.
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5.1.3 Verification of the titin-DsRed protein and its localization within the
sarcomere
Since introducing of the DsRed coding sequence altered the titin gene, we had to exclude that
expression of titin-DsRed results in an obvious phenotype regarding viability and fertility.
Therefore, offspring from breeding of heterozygous male and female mice was investigated for
the distribution of the genotype. We found normal Mendelian ratios of 23.25% (n=10) wild-type,
53.5% (n=23) heterozygous knockin, and 23.25% (n=10) homozygous knockin mice examining
in total 43 animals. Also the distribution of 46.5% (n=20) males and 53.5% (n=23) females were
as expected and mice of all genotypes had a normal lifespan. Figure 12 shows that the body
weight with ~24.5 g and the ratio of heart weight to body weight (corrected heart weight) with
~4.8 mg/g for wild-type (+/+), heterozygous (R/+) and homozygous (R/R) female mice were
unchanged. These data indicate that the integration of the DsRed into the titin protein did not
affect apparently the structure and function of the heart and skeletal muscle.
Figure 12: Titin-DsRed knockin mice did not have an obvious phenotype. Heterozygous (+/R; n=6) and
homozygous (R/R; n=5) female animals had an equal body weight (A) as well as heart weight to body weight ratio
(B) compared to the littermate wild-type control (+/+; n=7; mean with SEM; One-way ANOVA P>0.5 ).
TiEx28DsRed mice did not have an obvious phenotype so that it was possible to further
investigate the mouse model on protein level. Therefore, we used a SDS-agarose gel to separate
the titin-isoforms and Western blotting (Figure 13) using samples of the striated muscles heart,
quadriceps, and soleus, in which titin is well studied.
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Figure 13: Detection of titin-DsRed in heart, quadriceps, and soleus. A) Coomassie stained SDS-agarose gel
shows separation of the T1 N2B (2970 kDa) and N2BA (3300 kDa) isoforms of the heart, the two N2A isoforms of
the quadriceps and the long N2A isoform (3700 kDa) in the soleus as well as the T2 degradation product
(~2000 kDa) in samples from wild-type (+/+; n=3), heterozygous (R/+; n=3) and homozygous (R/R; n=3) mice. B)
Western blot using a DsRed antibody verified the presence of DsRed in T1 full-length titin of heterozygous (n=3)
and homozygous knockin mice (n=3). The M8/M9 antibody used in Western blot detected T1 and T2 titin
demonstrating the translation of titin’s C-terminus in striated muscle samples of all genotypes (n=3).
The SDS-agarose gel shows that it was possible to separate the T1 N2B (2970 kDa) and N2BA
(3300 kDa) isoforms of the heart and the two T1 N2A isoforms of the quadriceps as well as the
T2 degradation products (Figure 13A). In the soleus, the longest T1 N2A isoform (3700 kDa)
was observed. All titin isoforms and the corresponding smaller T2 degradation products
(~2000 kDa) were present in wild-type (+/+), heterozygous (R/+), and homozygous (R/R) mice
demonstrating that the DsRed did not affect the expression and translation of full length titin in
striated muscle. Furthermore, the existence of the same amount of titin isoforms in the three
genotypes in the heart, quadriceps, and soleus indicates that the DsRed insertion did not affect
protein stability. Since the DsRed has a calculated molecular weight of 26.8 kDa, which is small
compared to titin isoforms of up the 3700 kDa in striated muscle, it was not possible to separate
in heterozygous knockin mice the titin-DsRed from the wild-type titin molecules. To ensure, that
the DsRed was present within titin, Western blot was performed using a DsRed antibody (Figure
13B). We detected a strong signal from the T1 isoforms in the heart, quadriceps, and soleus
samples of homozygous mice and a weaker signal in the samples of heterozygous mice because
homozygous knockin mice carry only the titin-DsRed and the heterozygous knockin mice
expressed titin-DsRed as well as wild-type titin. The titin isoforms in the samples from wild-
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type littermate controls served as a negative control. Figure 13B confirms with the help of the
M8/M9 antibody, which is reacting against titin’s C-terminal region, that full length titin was
translated in wild-type, heterozygous, and homozygous mice. In contrast to the DsRed-antibody,
which did not detect the degradation product T2, the M8/M9 antibody identified T1 and T2.
As intended by the targeting strategy, DsRed was expressed and translated within full length titin
in knockin mice of the TiEx28DsRed strain. To investigate the DsRed localization and its
influence on titin’s integration into the sarcomere, fluorescence microscopy of cryosections was
performed using quadriceps as an example of skeletal muscle and heart (Figure 14). It was
possible to monitor the fluorescence emitted by the DsRed (red) in heterozygous (R/+) and
homozygous (R/R) knockin mice in both heart (A) and quadriceps (B). A weaker signal was
detected in heterozygous mice because they carry titin-DsRed as well as wild-type titin. No
signal was detected in control wild-type mice (+/+). The DsRed fluorescence appeared as
striated pattern indicating that the incorporation of titin into the sarcomere and thereby also
sarcomere assembly was normal. The localization of the DsRed close to titin’s N-terminus was
confirmed using an M8/M9-antibody (green) that marks the M-band. This immunostaining led
to a striated pattern for all genotypes in heart and quadriceps due to the highly ordered
arrangement of the sarcomeres in striated muscle. Overlay with the DsRed fluorescence
(overlay) shows an alternating striated pattern. This demonstrates the constant series of Z-discs
and M-bands in the striated muscle of heterozygous and homozygous knockin mice confirming
the presence of DsRed at the Z-disc. Thus, titin-DsRed integrated into the Z-disc and M-band so
that the sarcomeric structure was not influenced by the insertion of the DsRed into the titin’s Z/I-
junction. The targeting strategy was successful and we were able to establish the new titin-
DsRed knockin mouse model TiEx28DsRed, which will help to visualize and analyze titin in
smooth muscle.
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Figure 14: The DsRed located close to titin’s N-terminus did not influence the sarcomeric structure.
Fluorescence microscopy of (A) heart and (B) quadriceps cryosections obtained from wild-type (+/+; n=3),
heterozygous (R/+; n=3) and homozygous (R/R; n=3) mice. Fluorescence was emitted by the DsRed (red) in
heterozygous and homozygous knockin animals and appeared as striated pattern demonstrating that the sarcomere
assembly was normal. Immunostaining with an antibody against the immunoglobulin domains M8 and M9 at titin’s
C-terminus marking the M-band resulted in a striated pattern for all genotypes (green). The overlay of the M8/M9
staining with the DsRed fluorescence shows an alternating striated pattern (overlay) demonstrating a constant series
of Z-discs and M-bands in striated muscle of heterozygous and homozygous mice. This indicates the presence of
the DsRed close to titin’s N- terminus. Bar: 5 m.
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5.2 Characterization of the smooth muscle specific knockout of
titin’s kinase region
5.2.1 Establishing the smooth muscle specific mouse model
For analysis of the titin’s kinase region in smooth muscle, the conditional knockout mouse
model TiMEx1-2 (Gotthardt et al., 2003) was used, in which titin’s M-band exons 1 and 2 that
correspond to titin’s exons 358 and 359, are flanked by two lox-sites (Figure 15 floxed allele).
The conditional mouse model was bred with a transgenic mouse strain (SMMHCcre), which
expresses the cre-recombinase under the control of the smooth muscle myosin heavy chain
promoter, mediating the deletion of titin’s M-band exons 1 and 2 by recombination of the lox-
sites in the generated SMcTiMEx1-2 mouse strain (Figure 15 deleted allele).
Figure 15: Strategy to delete titin’s kinase region in smooth muscle. Exon/intron (boxes/lines) structure of the
titin gene close to its 3’end as present in the conditional knockout model TiMEx1-2. Titin’s M-band exons 1 and 2
(titin exons 358 and 359) encoding the kinase region, which are flanked by two lox-sites (floxed allele), are deleted
(deleted allele) by breeding with transgene animals expressing the cre-recombinase under the control of the smooth
muscle myosin heavy chain promoter (SMMHCcre) leading to the SMcTiMEx1-2 mouse strain. Primers for
genotyping the titin alleles are indicated as arrows. Modified from Gotthardt et al., 2003.
Our aim was to study the effect of the deletion of titin’s kinase region in smooth muscle on
uterus, because pregnancy involves contraction as well as tissue remodeling. It has been shown,
that the SMMHC cre-recombinase is present from E12.5 in smooth muscle (Xin et al., 2002),
but its recombination activity in uterine tissue has not been confirmed. Therefore, SMMHCcre
mice were crossed with GtROSA26 reporter mice to monitor cre-recombinase activity in the
generated SMcGtROSA26 strain. Figure 16 shows the lacZ staining of cryosections obtained
from uteri in the estrous stage of 90 d old adult females. The sections were counterstained with
eosin so that the cytoplasm and connective tissue appears red.
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Figure 16: Activity of the SMMHC cre-recombinase in uterine smooth muscle. LacZ staining (blue)
counterstained with eosin (red) of uterine cross sections shows that the SMMHCcre was active leading to
recombination in circular and longitudinal smooth muscle cells in the uterus of animals that contain the lacZ
reporter gene as well as the cre-recombinase transgene (lacZ+ cre+; n=3). The negative controls, either the lacZ
reporter gene (lacZ+, cre-; n=3) or the cre-recombinase transgene (lacZ-, cre+; n=3), did not give a signal in the
LacZ staining. cSMCs: circular smooth muscle cells; ge: glandular epithelium; l: lumen; le: luminal epithelium;
lSMCs: longitudinal smooth muscle cells; myo: myometrium; s: stroma ; bar: 100 m.
In animals that carried the lacZ reporter gene (lacZ+) as well as the cre-recombinase transgene
(cre+) recombination mediated by the cre-recombinase has occurred in myometrial circular and
longitudinal smooth muscle cells. This cre-recombinase activity led to lacZ expression and the
presence of the -galactosidase, which is visible by the blue lacZ staining. The negative staining
controls of mice containing either the lacZ reporter gene (lacZ+, cre-) or the cre-recombinase
transgene (lacZ-, cre+) did not show lacZ expression indicating that endogenous -galactosidase
is not sufficient for recombination and the cre-recombinase alone cannot cause lacZ staining.
Since we could show that the cre-recombinase under the control of the smooth muscle myosin
heavy chain promoter is active in the uterine smooth muscle, we investigated recombination in
uterus and tail of SMcTiMEx1-2 and SMMHCcre animals in estrous stage by PCR (Figure
17A).
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Figure 17: Recombination by the SMMHC cre-recombinase in uterine smooth muscle and the germline. A)
PCR-analysis of uterine and tail tissues obtained from control (CON), heterozygous (HET), and knockout (KO)
mice of the SMcTiMEx1-2 strain and animals of the SMMHCcre strain containing the transgene (Cre+) in estrous
stage. The tissues were genotyped for the presence of the floxed titin allele (lox) or the corresponding wild-type
allele (WT) and the cre transgene (cre). Recombination (rec) has occurred not only in the uterus of knockout and
heterozygous mice as expected, but also in the uterus of control mice and in tails of control and knockout mice
indicating smooth muscle but also germline activity of the cre-recombinase. Primers to obtain WT, lox, and rec
signals are given in Figure 15. B) Real-Time PCR analysis revealed the expression of titin’s exons encoding parts of
the N-terminus (ZEx3), N2A region (N2A) and C-terminus (MEx6) in control (n=3), heterozygous (n=3), and
knockout (n=3) mice as well as the lack of mRNA encoding titin’s kinase region (MEx1-2) in knockout animals
(mean with SEM; Two-way ANOVA P<0.001 ***). The mRNA levels were normalized to 18S RNA and expression
is displayed as fold change relative to the control.
To confirm the genotype of control, heterozygous, and knockout mice of the SMcTiMEx1-2
strain and of SMMHCcre animals containing the transgene (Cre+), tail and uterine tissues were
first analyzed for the presence of the floxed titin allele as well as the cre transgene. As expected
for both tissues, control and knockout mice carried homozygous the floxed allele (lox),
heterozygous mice had a floxed as well as a wild-type allele (WT), and Cre+ control mice had
only the titin wild-type allele. The cre transgene (cre) was present in the Cre+ control,
heterozygous, and knockout mice. DNA of control mice did not lead to a signal in the cre-PCR.
The PCR detecting the recombination of the floxed titin allele (rec) revealed that recombination
occurred in uteri of knockout, heterozygous, and unpredicted also in control mice, but not in
Cre+ control animals. Additionally, control and homozygous mice were recombined in the tail.
Since recombination was observable in uteri of control animals and also in tails of control and
knockout mice, we concluded that beside the smooth muscle recombination, which is shown by
the recombination in heterozygous animals and in Figure 16, also unspecific recombination has
happened independent of the presence of the cre-recombinase. This finding was confirmed by de
Lange et al. (de Lange et al., 2008), who described the germline activity of the SMMHC
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promoter. They showed that the activity of the cre-recombinase in male germ cells is irrespective
of the cre-genotype and ends prior to fertilization.
The amount of titin mRNA after deletion of titin’s M-band exons 1 and 2 in non-pregnant uterus
was quantified by Real-Time PCR (Figure 17B). It revealed the expression of titin regions
including the N2A region (N2A) as well as the N- and C-terminus represented by the Z-disc
exon 3 (ZEx3) and the M-band exon 6 (MEx6) in uterus of control, heterozygous, and knockout
mice of the SMcTiMEx1-2 strain. In opposition to control and heterozygous mice, the mRNA of
titin’s M-band exons 1 and 2 (MEx1-2) could not be detected anymore in knockout animals.
Nevertheless, the expression of titin’s C-terminus was not affected by the deletion of the M-band
exons 1 and 2 coding for the kinase region in these females.
5.2.2 Titin’s kinase region is essential for embryo implantation
5.2.2.1 Normal preimplantational embryo development in titin’s kinase region
deficient mice
We were able to efficiently delete titin’s M-band exons 1 and 2 encoding the kinase region in
knockout mice. Knockout animals did not have an obvious phenotype so that we generated a
suitable mouse model to address a smooth muscle function of titin’s kinase region during
pregnancy. To distinguish effects on non-pregnant uterus and preimplantational embryonic
development from effects during embryo implantation, we investigated the estrous cycle length
of non pregnant uteri and the number of E3.5 blastocysts (Figure 18).
Figure 18: Normal estrous cycle length and number of blastocysts in knockout females. A) The days (d) of
estrous cycle length did not differ between control (CON; n=7) and knockout animals (KO; n=7; mean with SEM;
T-test P>0.05). B) The number of blastocysts at E3.5 recovered from control (n=23) and knockout animals (n=19)
was the same in both groups (mean with SEM; T-test P>0.05).
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The estrous cycle did not vary between knockout and control mice (~10.0 d and ~10.4 d,
respectively; Figure 18A). Furthermore, metestrus, diestrus, proestrus, and estrus were clearly
distinguishable in control and knockout females including the typical presence of leucocytes and
epithelial cells. Hence we concluded that the deletion of titin’s kinase region did not affect the
recurring changes of the non pregnant uterus in response to ovarian hormones so that we were
able to set up timed matings to obtain pregnant females. The amount of blastocysts at E3.5 was
the same for control and knockout mice (~8; Figure 18B). There was also no difference in
morphology of blastocysts recovered from control and knockout uteri. These data demonstrated
normal ovulation, fertilization, transport of the developing embryo through the oviduct, and
blastocyst development.
5.2.2.2 Impaired implantation affects embryo positioning, number of implantation
sites, and uterine maturation
Embryo implantation studies were done at E4.5 and E5.5 to investigate the implantation time
and positioning of the embryos within the uterus. Figure 19 shows that the loss of titin’s kinase
region led to clear phenotypic changes as visualized by Evans Blue staining.
Figure 19: Impaired embryo implantation in knockout females. Evans Blue staining at E4.5 and E5.5 shows on-
time implantation that is impaired in knockout (KO) uteri resulting in a reduced number of implantation sites (IS <
5, blue), mislocalization (white arrows indicate lack of implantation sites), and crowding (black arrows) compared
to the equally distributed implantation sites in control (CON) mice. The percent distribution as well as the total
number of investigated animals is indicated. Bar: 1 cm.
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Embryo implanted on-time in knockout females because implantation sites (IS) were already
detectable at E4.5 as in control mice. The implantation sites were equally distributed on both
uterine horns and evenly spaced in ~90% of the control uteri at E4.5 and E5.5. In contrast, only
~39% of the knockout mice had a normal implantation pattern at either time point. We classified
the uteri with impaired implantation into three groups with consistent distribution at E4.5 and
E5.5. Knockout uteri showed in 22.7% less than five implantation sites and in 13.6% crowding
of two or more implantation sites, whereas this clustering did not occur in a special segment of
the uterus. The positioning of the implantation sites were considered to be mislocalized, if more
than triple implantation sites were found in one uterine horn compared to the other. This was the
case in ~24% of knockout animals. Rarely, knockout uteri exhibited multiple defects.
We investigated the number of implantation sites at E4.5 and E5.5 of control and titin kinase
deficient knockout mice in more detail (Figure 20A).
Figure 20: Reduced number of implantation sites in knockout uteri implies a defect during the penetration
phase. A) The number of implantation sites at E4.5 and E5.5 varied more in knockout mice (KO; E4.5 n=22; E5.5
n=22) compared to control animals (CON; E4.5 n=19; E5.5 n=23) and was additionally significant reduced by
~26% (mean with SEM; Two-way ANOVA P<0.01 **). B) In one knockout uterus it was possible to observe
embryos that have attached but did not develop further as shown by lack in Evans Blue staining at E5.5 (white
arrows). Bar: 1 cm.
At E4.5 and E5.5, the number of implantation sites varied more in knockout animals compared
to control mice and was additionally significantly reduced by ~26%. Since comparable numbers
of blastocysts were available for implantation in control and knockout uteri and the diminished
amount of implantation sites in knockout uteri was identical at E4.5 and E5.5, we concluded that
the loss of titin’s kinase region led to defects during the penetration phase of embryo
implantation, whereas apposition and attachment were not affected. This finding was confirmed
by one knockout uterus, in which four implantation sites have not further developed (Figure
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20B). It was also not possible to get blastocysts from E4.5 uteri that showed a reduced number
of implantation sites indicating proper embryo attachment.
The change of the uterine wet weight in response to embryo implantation was determined as an
index for vascularization and decidualization, which occur during the penetration phase. The
uterus weight of knockout and control animals at E3.5, E4.5, and E5.5 was measured and
normalized to the body weight. E3.5 was confirmed by the presence of blastocysts and only uteri
with ten implantation sites were taken into account for E4.5 and E5.5 (Figure 21).
Figure 21: Delayed development of knockout uteri during the penetration phase. A) Between E3.5 to E5.5 the
uterus weight to body weight (uw/bw) ratio of knockout (KO) and control (CON) animals increased continuously.
There was no difference between knockout and control animals at E3.5 (KO n=10; CON n=11) and E4.5 (KO n=4;
CON n=4), but knockout uteri showed a decreased uterus weight to body weight ratio at E5.5 (KO n=6; CON n=6;
mean with SEM; Two-way ANOVA P<0.01 **). E3.5 was determined by the presence of blastocysts and for E4.5
and 5.5 only uteri with 10 implantation sites were used for evaluation. B) The delayed development of the knockout
uteri was independent of the body weight (bw), which did not differ between control and knockout mice at E3.5,
E4.5, and E5.5 (mean with SEM; Two-way ANOVA P>0.05).
The uterus weight to body weight (uw/bw) ratio of knockout and control animals increased
steadily from E3.5 to E5.5 (Figure 21A). In contrast to E3.5 and E4.5, knockout uteri had a
decreased uterus weight to body weight ratio at E5.5 in comparison to control females. This
reduction in the uterus weight to body weight ratio was independent of the body weight that did
not vary between the two groups and the investigated time points (Figure 21B). Since the
increase of the uterus weight in the milligram range cannot be due to the developing embryo
during early pregnancy, we could show that the absence of the titin kinase region in smooth
muscle led to delayed uterine development in the penetration phase of embryo implantation.
Additionally, the heart weight to body weight ratio was determined using the same set of
animals (supplement Figure 31). There was no difference between control and knockout animals
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indicating that the deletion of titin’s kinase region in vascular smooth muscle cells did not cause
an obvious phenotype.
To exclude, that the presence of the cre-recombinase in smooth muscle of knockout animals
resulted in the implantation defects, embryo implantation studies were also conducted with
heterozygous and Cre+ control animals that contained the cre-recombinase (Figure 22). E5.5
was investigated because at this time point knockout females had both impaired implantation
and delayed uterine development. Only uteri with ten implantation sites were used for
evaluation.
Figure 22: The phenotype observed in the knockout females was due to the deletion of titin’s kinase region.
A) Although containing the cre-recombinase, Cre+ control (Cre+) and heterozygous (HET) mice had normal
embryo spacing and distribution as well as more than four implantation sites. The percent distribution and the total
number of investigated animals is given. Bar: 1 cm. B) At E5.5, the uterus weight to body weight (uw/bw) ratio of
Cre+ control and heterozygous animals was similar to the control animals (CON). The knockout (KO) uterus weight
to body weight ratio was reduced compared to the other three groups (CON n=6; HET n=5; KO n=6; Cre+ n=3;
mean with SEM; one-way ANOVA P<0.05 * and P<0.001 ***). Only uteri with 10 implantation sites were taken
into account.
As shown by Evans Blue staining in Figure 22A, 92% of the Cre+ control mice and 78% of the
heterozygous mice had a normal implantation including even embryo spacing and a normal
number of implantation sites as the control mice (Figure 19). The uterus weight to body weight
ratio of Cre+ control mice and heterozygous mice was similar to the control animals, which
were cre-recombinase negative. In contrast, the uterus weight to body weight ratio at E5.5 of
knockout animals was reduced compared to littermate control animals as well as to Cre+ control
and littermate heterozygous mice (Figure 22B). This effect was independent of the body weight
(supplement Figure 31B). Hence we concluded that the phenotype observed in knockout females
was not due to the cre-recombinase, but caused by the deletion of titin’s kinase region.
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5.2.3 Loss of titin’s kinase region influences arachidonic acid metabolism
It has been shown that mice with null mutation for Alox15 that encodes the leukocyte 12/15-
lipoxygenase (L-12/15-LOX) have a reduced number of embryo implantation sites (Li et al.,
2004) similar to the phenotype we observed in the knockout animals lacking titin’s kinase region
(Figure 20). Hence we investigated the mRNA level (Figure 23A) of Alox15 as well as of the
genes Pparg, Creb1, and Mapk13 encoding the peroxisome proliferator-activated receptor
(PPAR), the cAMP responsive element binding protein 1 (CREB1), and the mitogen-activated
protein kinase 13 (MAPK13), which have been implicated to be possible downstream targets of
metabolites derived from the L-12/15-LOX (Reddy et al., 2002; Li et al., 2004). Uterine tissue
of E3.5 pregnant mice was used because previous work has demonstrated that Alox15 has the
highest expression level at E3.5 (Li et al., 2004).
Figure 23: Deletion of titin’s kinase region affected arachidonic metabolism. A) Real-Time PCR analysis
revealed the upregulation of Alox15 mRNA as well Mapk13 mRNA in knockout (KO; n=3) compared to control
(CON; n=3) animals using E3.5 uterine tissue. The genes Pparg and Creb1 encoding two other possible
downstream targets of the L-12/15-LOX, the PPAR and the CREB1, were unchanged on mRNA level. The mRNA
levels were normalized to 18S RNA and expression is displayed as fold change relative to the control (mean with
SEM; T-test comparing CON and KO of one primer/probe set P<0.05 * and P<0.01 **). B) Compared to control
animals (n=12), the uterine levels of 12-HETE, which is produced from arachidonic acid by the L-12/15-LOX, were
reduced by ~20% in E3.5 pregnant knockout females (n=13). The metabolite concentration is displayed relative to
the total protein amount in the sample (mean with SEM; T-test P<0.05 *).
Figure 23A shows mRNA upregulation of Alox15 and Mapk13 in knockout animals compared to
control animals. There was no significant difference in the uterine mRNA level of Pparg and
Creb1 between both groups. Since the L-12/15-LOX catalyzes among others the formation of
12-HETE from arachidonic acid that is important for embryo implantation (Li et al., 2004),
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uterine 12-HETE levels of E3.5 pregnant mice were measured (Figure 23B). The uterine 12-
HETE levels in knockout animals were ~20% less in comparison to control animals.
5.2.4 Aberrant angiogenesis in knockout animals lacking titin’s kinase
region
The adaption of the uterus to enable implantation of the embryo requires changes in vascular
permeability and angiogenesis. Since the SMcTiMEx1-2 knockout females had a reduced
uterine wet weight in response to embryo implantation (Figure 21), which is a measurement for
vascularization and decidualization, we investigated angiogenesis in vivo in female and male
mice (Figure 24A). The angiogenesis was induced by VEGF because this angiogenic growth
factor has been shown to be a key regulator for angiogenesis (Carmeliet et al., 1996; Fong et al.,
1995) and also to be necessary for implantation to occur (Rockwell et al., 2002).
Figure 24: Altered angiogenesis and PDGF-B serum levels in knockout mice. A) VEGF-induced angiogenesis
in vivo was ~60% reduced in knockout (KO) females (n=10) and ~54% in knockout males (n=8) in comparison to
the corresponding control group (CON females n=10; CON males n=10). The vascularization was measured by
assessing the hemoglobin content of matrigel plugs, which is expressed as blood volume content within the matrigel
plug (mean with SEM; Two-way ANOVA P<0.05 *). B) Multiplex bead immunoassay to determine cytokines in
pregnant E3.5 mice revealed a ~34.5% increase in the PDGF-B serum levels of knockout animals (n=13) in
comparison to control animals (n=14; mean with SEM; T-test P<0.05 *). The same amount of VEGF was present in
the serum of knockout (n=16) and control (n=16) animals (mean with SEM; T-test P>0.05). The metabolite
concentration is illustrated per serum volume.
In vivo blood vessel formation, which was measured as blood volume content within matrigel
implants, was reduced ~60% in knockout females and ~54% in knockout males compared to the
control mice (Figure 24A). Hence we concluded that the deletion of titin’s kinase region in
smooth muscle cells resulted in an angiogenesis defect, which was independent of the gender. To
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correlate the defects in angiogenesis and embryo implantation with changes in cytokine levels,
the content of FGF basic, G-CSF, IL-1β, IL-6, IL-15, IP-10, LIF, MCP-1, PDGF-B, TNF-α, and
VEGF was determined in serum and uterine tissue of E3.5 pregnant mice. We found a ~34.5%
increase of the growth factor PDGF-B in sera obtained from knockout mice compared to the
control group (Figure 24B). Serum VEGF levels were unchanged indicating that impaired
angiogenesis in knockout animals was not due to a defect in responding to VEGF. Also the other
cytokines did not differ in the serum (supplement Figure 32) as well as in the uterine levels
(supplement Figure 33). It was not possible to detect FGF basic, LIF, Il-15, IP-10, and MCP-1 in
the serum and FGF basic, G-CSF, Il-15, PDGF-B, and VEGF in the uterine tissue.
Since PDGF-B is responsible for the migration of pericytes and vascular smooth muscle cells to
the site of angiogenesis (Lindahl et al., 1997; Hellström et al., 1999), we tested the ability of
smooth muscle cells that were isolated from the aorta of knockout and control animals to
migrate in response to 10-10 M PDGF-B (Figure 25A). The migration rate of smooth muscle cells
is indicated relative to the corresponding control without chemoattractant, which was set to be
100%.
Figure 25: Knockout smooth muscle cells did not migrate in response to PDGF-B. A) Migration study with
smooth muscle cells isolated from the aorta of knockout (KO; n=6) and control (CON; n=6) animals using 10-10 M
PDGF-B as chemoattractant revealed that knockout smooth muscle cells failed to migrate. In contrast, control
smooth muscle cells showed a ~69% increase in migration in response to PDGF-B. The migration rate is given
relative to the migration of the corresponding smooth muscle cells without chemoattractant, which was set to 100%
(mean with SEM; T-test P<0.05 *). B) Immunostaining using an antibody against smooth muscle actin (red) of
cultures used for the migration studies confirmed the presence of >90% knockout (n=6) and control (n=6) smooth
muscle cells. Nuclei of the cells were co-stained with DAPI (blue) to evaluate the amount of smooth muscle actin
positive cells. Bar: 100 m.
Knockout smooth muscle cells did not migrate in response to PDGF-B because they had the
same migration rate as knockout smooth muscle cells without chemoattractant (Figure 25A). On
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74
the contrary, smooth muscle cells obtained from control animals migrate by PDGF-B stimulation
~69% more compared to their basal migration rate. The presence of >90% smooth muscle cells
in cultures used for the migration study was confirmed by detecting the cell type restricted
intermediate filament smooth muscle actin (Figure 25B). Since knockout smooth muscle cells
did not migrate in comparison to control smooth muscle cells, we concluded that the loss of
titin’s kinase region in smooth muscle cells affects cell migration. We assume that this migration
defect led to impaired in vivo angiogenesis in knockout mice (Figure 24A).
5.2.5 Alteration of tissue remodeling in titin’s kinase region deficient
animals
5.2.5.1 Expression of Murf1 and Fhl2 depends on titin’s kinase region
The homozygous deletion of titin’s M-band exons 1 and 2 has caused the loss of binding sites
for MuRF1, MuRF2, T-cap, NBR1, p62, calmodulin (CALM1), FHL2, and myomesin (Myom1)
in knockout animals (Figure 29). To elucidate, which genes encoding these M-band binding
proteins are expressed in smooth muscle and contribute to the phenotype, expression analysis
was performed using uterine tissue of E3.5 pregnant mice (Figure 26).
Figure 26: Altered expression of Murf1 and Fhl2 in knockout uteri. Real-Time PCR was performed to analyze
the mRNA level of genes encoding proteins that bind to titin’s kinase region, which is deleted in the knockout
animals. Murf1 expression was upregulated in knockout animals (KO; n=3) compared to control mice (CON; n=3)
using uterine tissue from E3.5 pregnant females. Moreover, the transcription level of Fhl2 was reduced in knockout
uteri. The expression of Nbr1, p62, Calm1, and Myom1 was similar in both groups. The mRNA levels were
normalized to 18S RNA and expression is displayed as fold change relative to the control (mean with SEM; T-test
comparing CON and KO of one primer/probe set P<0.05 *).
The expression of Murf1 was upregulated in knockout animals in comparison to control mice
(Figure 26). Furthermore, the transcript level of Fhl2 was little but significant diminished in the
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75
uterine tissue of knockout mice. The expression of Nbr1, p62, Calm1, and Myom1 was
unaffected in knockout and control females. The mRNA of the titin M-band binding proteins T-
cap and MuRF2, which forms heterodimers with MuRF1, could not be detected in uterine tissue.
Since titin’s kinase region has been implicated as a regulator of contractile function by effecting
calcium handling (Peng et al., 2007), we also investigated the expression of the genes encoding
the calcium-related proteins PKC, PKC, S100A1, L-type calcium channel (CACNA1C),
calcium/calmodulin-dependent protein kinase II alpha (CAMK2A), and calbindin 1 (CALB1).
The expression level of these genes was unchanged in E3.5 pregnant uterine tissue (supplement
Figure 34).
5.2.5.2 The transcription level of Cox2 and Lpar3 is not affected
Reproductive failure including reduced litter size and implantation defects is frequently
associated with changes in expression of the genes Cox2 and Lpar3 encoding COX2 and the
lysophosphatic acid receptor LPA3 (Lim et al., 1997; Matsumoto et al., 2002; Ye et al., 2005). It
has also been shown that LPA3-mediated signaling regulates embryo spacing and implantation
timing (Hama et al., 2007; Ye et al., 2005). To investigate if the transcript level of Cox2 and
Lpar3 was affected upon the loss of titin kinase region, uteri of E3.5 pregnant control and
knockout animals were analyzed by Real-Time PCR (Figure 27).
Figure 27: Loss of titin’s kinase region did not alter the expression of Cox2 and Lpar3. The transcript level of
the genes Cox2 and Lpar3 encoding COX2 and LPA3, which are both known as regulators of embryo implantation,
were unchanged in knockout (KO; n=3) compared to control (CON; n=3) animals investigating E3.5 pregnant uteri.
The mRNA levels were normalized to 18S RNA and expression is displayed as fold change relative to the control
(mean with SEM; T-test comparing CON and KO of one primer/probe set P>0.05).
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76
As shown in Figure 27, the expression of Cox2 as well as Lpar3 was not significantly changed
in knockout animals compared to control mice.
5.2.5.3 Alterations of structural and ubiquitin-proteasome related proteins
Proteins related to the progression of the phenotype caused by titin’s kinase region deficiency
were investigated using 2D-gel electrophoresis. Uterine tissue of E3.5 pregnant knockout and
control animals was analyzed (supplement Figure 35). In total we have found 106 proteins that
were altered comparing knockout and control animals. 32 proteins were downregulated and 74
proteins were upregulated in the knockout uteri, of which 20 could be identified so far (Figure
28, supplement Table 11). ANXA2, CKB, HSPB1, PDIA4, PSMC2, TAGLN were reduced in
the knockout uteri and ARPC2, CAPG, CAR2, COL6A2, DNAJA1, ETFDH, FGA, HNRNPAB,
NDUFS3, PCX, SFRS3, TPI1, and VIM accumulated in the knockout uteri (Figure 28).
Depending on their function, these proteins belong to metabolism (CKB, ETFDH, NDUFS3,
PCX, TPI1) or the heat shock and proteasomal stress response (HSPB1, PDIA4, PSMC2,
DNAJA1), are related to the cytoskeleton (TAGLN, ARPC2, CAPG, VIM), or have other
functions (ANXA2, CAR2, COL6A2, FGA, HNRNPAB, SFRS3). Hence we concluded that loss
of the titin kinase region affected uterine tissue remodeling since structural proteins as well as
proteins of the ubiquitin-proteasome pathway were changed.
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77
Figure 28: Impaired heat shock response and altered cytoskeleton-related proteins in knockout uteri. 2D-gel
electrophoresis was performed to compare the proteins of E3.5 pregnant uteri of control (CON; green; n=6) and
knockout (KO; red; n=6) mice. The Delta2D software colorized the control protein spots green and the knockout
protein spots red leading in the dual view to yellow spots, if the same amount of protein is present in both groups
(T-test P<0.1 *; white arrows point the changed protein spots). Downregulation in knockout uteri is indicated by the
green spots in the dual view for the proteins annexin A2 (ANXA2), brain creatine kinase (CKB), heat shock
protein 1 (HSPB), protein disulfide-isomerase A4 (PDIA4), proteasome 26S subunit ATPase 2 (PSMC2), and
transgelin (TAGLN). Accumulation of proteins in knockout uteri is indicated by red spots in the dual view for the
proteins actin related protein 2/3 complex (ARPC2), capping protein (CAPG), carbonic anhydrase 2 (CAR2),
collagen type VI alpha 2 (COL6A2), DnaJ (Hsp40) homolog (DNAJA1), electron transferring flavoprotein
dehydrogenase (ETFDH), fibrinogen (FGA), heterogeneous nuclear ribonucleoprotein A/B (HNRNPAB), NADH
dehydrogenase (ubiquinone) Fe-S protein 3 (NDUFS3), pyruvate carboxylase (PCX), splicing factor
arginine/serine-rich 3 (SFRS3), triosephosphate isomerase 1 (TPI1), and vimentin (VIM).
Discussion
78
6 Discussion
Aim of this study was the functional and morphological characterization of titin in smooth
muscle. So far, only expression profiling, localization, and in vitro interaction studies for smooth
muscle titin and the titin-like protein smitin have been performed, but a role has not been
assigned (Kim and Keller, 2002; Chi et al., 2005, 2008; Labeit et al., 2006). Therefore, a smooth
muscle specific knockout of titin’s M-band exons 1 and 2 was used in our research to address a
function for titin in uterine smooth muscle. Furthermore, the new titin-DsRed knockin mouse
model TiEx28DsRed was established to facilitate the detection of smooth muscle titin.
In the TiEx28DsRed mouse strain, the coding sequence of the red fluorescence protein DsRed
was inserted into exon 28 of the titin gene (Figure 7). The advantage of the presence of a
fluorescence protein within titin is live cell imaging and the availability of commercial
antibodies against DsRed. In contrast, there are only few titin antibodies purchasable. Hence the
DsRed will help to detect smooth muscle titin, especially because the transcript level of smooth
muscle titin is ~100 fold less than in skeletal muscle (Labeit et al., 2006) and therefore its
verification on protein level limited. Furthermore, only the domain composition of one smooth
muscle isoform is proposed based on expression analysis of human aorta, bladder, carotid, and
stomach (Labeit et al., 2006). Since titin in striated muscle is alternatively spliced during
embryonic development as well as in different tissues (Labeit and Kolmerer, 1995; Cazorla et
al., 2000; Freiburg et al., 2000; Bang et al., 2001a; Sorimachi et al., 1997), it is also likely that
different smooth muscle titin isoforms exist to determine muscle elasticity. Hence it is not
known, if the antibodies used in the previous smooth muscle titin studies will also react against
other tissue, such as uterus. The only study that has been done in mammals on protein level
detected smooth muscle titin in porcine aorta and stomach by Western blotting as well as in
cultured human aortic smooth muscle cells and in the tunica media of bovine aorta by
immunofluorescence (Labeit et al., 2006). Smitin, the titin-like protein found in chicken gizzard
muscle has been localized in the smooth muscle contractile apparatus as well as detected by
Western blotting (Kim and Keller, 2002). So far, smooth muscle titin has not been detected on
protein level in murine tissue or cells, which is required to gain more confidence to study
smooth muscle titin by a loss of function approach with available titin knockout mouse models.
A publication of Chi and colleagues have demonstrated by RT-PCR the expression of titin’s
exon 28 in human carotid artery, bladder, and uterus (Chi et al., 2008). Since our titin-DsRed
knockin mice carry the coding sequence of the DsRed within exon 28, the TiEx28DsRed mouse
Discussion
79
strain will help to conclusively show the existence of smooth muscle titin and its localization
within the contractile unit to incorporate titin in the model of smooth muscle structure.
After successful cloning of the targeting vector DsRedKI (Figure 8), the efficiency of its
homologous recombination into the titin locus of the ES cells was 1:192 (Figure 9). The
efficiency depends on the length of homologous sequence and the amount of repetitive sequence
that is present within the long arm of the targeting vector (Hasty et al., 1991). Furthermore,
isogenicity of the targeting vector to the ES cell line used alters gene targeting frequency (Deng
and Capecchi, 1992; van Deursen and Wieringa, 1992). As homologous recombination occurs
during the S-phase of the cell cycle, also cell cycle rates of the ES cell lines are important (Wong
and Capecchi, 1987; Udy et al., 1997). Accordingly, Udy and colleagues have found efficiencies
from 1:50 to 1:1033 with the same targeting construct depending on the ES cell lines used. The
achieved homologous recombination frequency of the DsRedKI into the titin gene was within
the expected range at the upper level of efficiency. Nevertheless, the ES cell clone E3 did not
incorporate the full targeting vector, as it was not possible to detect the DsRed in ES cells
differentiated into cardiomyocytes (Figure 10). Nevertheless, the neomycin resistance cassette
was incorporated into the titin locus of the ES cell clone E3 because this clone survived the
selection with the antibiotic G418 and was positive in the ES cell genotyping PCR (Figure 9).
This PCR did not detect the lack of the DsRed coding sequence since the forward primer binds
within the neomycin resistance cassette and the reverse primer outside of the targeting in front of
its short arm within the titin gene (Figure 7). For the ES cell clone F5, the integration of the
targeting vector DsRedKI into the titin locus was verified by PCR (Figure 9) as well as the
presence of the DsRed within titin and its proper localization within the sarcomere on protein
level (Figure 10). This clone was used to generate chimeras, in which ES cells contributed to the
germ line cells so that the DsRed-targeted titin gene was passed to the next generation. The
neomycin resistance cassette was removed with the help of the FLP-recombinase in animals
carrying heterozygous the targeted allele leaving only a FRT-site in the intron sequence between
exon 28 and 29 (Figure 7). These mice were used to generate the mouse colony of the
TiEx28DsRed strain (Figure 11) to exclude that knockin animals get a distinct phenotype caused
by the neomycin resistance cassette as reported previously (Kaul et al., 2000).
In heart and skeletal muscle titin is alternatively spliced in particular the PEVK and N2B region
as well as in the Ig domain regions of the I-band region (Labeit and Kolmerer, 1995; Cazorla et
al., 2000; Freiburg et al., 2000; Bang et al., 2001a). Furthermore, the short novex 3 isoform does
not contain titin’s A-band and M-band region as well as large parts of the I-Band region because
it has an alternative titin C-terminus (Bang et al., 2001a). Therefore the DsRed should be
Discussion
80
inserted close to titin’s N-terminus to be present in all titin splice isoforms (Figure 4). Splicing
also varies the number of Z-repeats at titin’s central Z-disc (Gautel et al., 1996; Sorimachi et al.,
1997). In contrast to rabbit uterus, in which the entire Z-repeat region is missing although titin
could be detected (Sorimachi et al., 1997), it has been shown that part of the Z-repeat region is
present in human smooth muscle including uterus (Chi et al., 2008). In addition, the insertion of
DsReds within titin’s Z-disc region could interfere titin signaling and the integration of titin’s N-
terminus into the Z-disc of the sarcomere by disturbing binding sites for sANK1 (Kontrogianni-
Konstantopoulos and Bloch, 2003), T-cap (Gregorio et al., 1998; Mues et al., 1998), -actinin
(Ohtsuka et al., 1997a, b; Gregorio et al., 1998), or obscurin (Bang et al., 2001a; Young et al.,
2001). The DsRed within titin’s Z-disc region could also cause a thicker Z-disc because a longer
titin molecule would displace the positions of binding sites for Z-disc proteins, such as -actinin
that links titin to the thin filament system (Ohtsuka et al., 1997a, b; Gregorio et al., 1998).
Changing the thickness of the Z-disc could influence the mechanical properties of the sarcomere
because the width of the Z-band is considered to be a fundamental property of the muscle fiber
type (Rowe, 1973; Yamaguchi et al., 1985) and correlates with the number of titin Z-repeats
(Sorimachi et al., 1997; Peckham et al., 1997). Furthermore, a thicker Z-disc could change the
stability of the sarcomere as well as sterically disturb possible interactions of titin binding
partners on opposite titin molecules that overlap in the Z-disc (Knupp et al., 2002). Hence titin’s
exon 28 was chosen for insertion of the DsRed coding sequence because it is located at titin’s Z-
disc to I-band junction, in which no protein binding sites are known. It encodes unique
sequences and Ig domains (Bang et al., 2001a). To not disrupt titin’s domain structure, the
DsRed coding sequence was inserted in front of the Ig domain I1 of titin’s I-band segment
preserving the reading frame (Figure 7).
Heterozygous as well as homozygous DsRed knockin mice had the same lifespan as wild-type
littermates as well as normal fertility and offspring at the expected Mendelian ratios. The
viability of knockin animals demonstrated that the DsRed did not affect the integration of the
titin-DsRed into the Z-disc of the sarcomere. Otherwise, homozygous knockin mice would be
embryonic lethal comparable to a titin M-band deficient mouse model, in which the lack of
titin’s integration into the M-band led to sarcomere disassembly and death at E11 (Weinert et al.,
2006). Proper sarcomere assembly was also confirmed by immunofluorescence studies of
quadriceps as example of skeletal muscle and heart, which show an alternating striated pattern
by DsRed emitted fluorescence and M8/M9 antibody staining (Figure 14). The integration of the
DsRed into titin did not affect its fluorescence properties as it was possible to monitor the
fluorescence emitted by the DsRed in heterozygous and homozygous knockin mice. In
Discussion
81
cardiomyocytes that we obtained by differentiating ES cells, the DsRed was only detected with
the help of an antibody (Figure 10) because the newly formed cardiomyocytes were not highly
packed with myofibrils so that the fluorescence was under the detection limit.
The body weight and heart weight to body weight ratio of TiEx28DsRed animals was
determined to investigate if expression of the titin-DsRed causes atrophy or hypertrophy of the
heart as it has been shown by several groups that mutations within the titin gene cause
cardiomyopathy (Radke et al., 2007; Granzier et al., 2009; Itoh-Satoh et al., 2002; Peng et al.,
2006). The body weight as well as heart weight to body weight ratio of heterozygous and
homozygous knockin mice was unchanged compared to the wild-type animals (Figure 12)
indicating that the DsRed within titin did not affect its signaling function leading to atrophy or
hypertrophy.
The titin-DsRed mouse model was further analyzed using heart, quadriceps, and soleus muscle.
Coomassie staining of wild-type, heterozygous, and homozygous TiEx28DsRed samples (Figure
13A) verified the presence of the N2BA and the shorter N2B isoform in heart, whereas the N2B
isoform was predominantly present consistent with previous studies (Neagoe et al., 2002).
Furthermore, the two N2A isoforms of quadriceps and the longest N2A isoform of soleus were
detected in the adult mice independent of the genotype. This indicates normal postnatal titin
splicing of skeletal muscle because after birth all muscle types initially express a single titin
isoform that gradually decreases in size, whereas a second titin isoform appears in quadriceps of
adult mice (Ottenheijm et al., 2009b). Using an antibody against DsRed, the presence of the
fluorescence protein was confirmed in heterozygous and homozygous mice. The intact full
length titin T1 but not in its break down product T2 was detected (Figure 13B). T2 constitutes
titin’s A-band and C-terminal region because it is formed by cleavage of T1 titin within the I-
band (Matsuura et al., 1991; Ohtsuka et al., 1992) so that the DsRed at titin’s N-terminal part
was cut off. In contrast, the M8/M9 antibody that is specific for titin’s C-terminal Ig domains
M8 and M9 reacted with T1 and T2 of heart, quadriceps, and soleus samples from wild-type,
heterozygous, and homozygous TiEx28DsRed mice (Figure 13B). Hence the DsRed did not
affect the expression and translation of titin.
Our titin-DsRed mouse model, which incorporates the coding sequence of the DsRed within
exon 28, was characterized in striated muscle, in which titin has been well studied. Mice that
carried heterozygous or homozygous the titin-DsRed did not have an obvious phenotype with
regard to fertility, heart weight to body weight ratio, and appropriate sarcomere assembly. DsRed
incorporated close to titin’s N-terminus at the Z/I-junction of the sarcomere was able to emit
fluorescence, but it was also possible to detect this protein with an antibody. Since it has been
Discussion
82
shown that exon 28 is expressed in human carotid artery, bladder, and uterus (Chi et al., 2008),
we established a suitable tool to visualize titin in embryonic and adult smooth muscle. In
particular we are interested in the localization of titin in the uterus to complement our analysis of
titin’s kinase region during early pregnancy.
Titin’s kinase region, which is encoded by titin’s M-band exons 1 and 2, has essential functions
in mature and in embryonic striated muscle as revealed by previous studies. Deletion of titin’s
kinase region during late embryonic development resulted in altered contractility of skeletal
muscle, disassembly of the sarcomere, and early death indicating a critical role for maintaining
the muscle structure (Gotthardt et al., 2003; Peng et al., 2006; Ottenheijm et al., 2009a).
Furthermore, results obtained by the loss of titin’s kinase region in adult mouse hearts suggest
that titin’s kinase region regulates contractile functions via calcium handling (Peng et al., 2007).
In the conventional knockout model of titin’s M-band exons 1 and 2, embryos died due to
mechanical instability of sarcomeres although the initial assembly of sarcomeres was unaffected
(Weinert et al., 2006). Since important functions for titin’s kinase region have been addressed for
striated muscle, we speculated that titin’s kinase region is also required in adult smooth muscle.
Moreover, titin’s kinase is highly conserved in vertebrate and invertebrates suggesting an
essential function in muscle tissue (Gautel et al., 1995). The importance of titin’s kinase in
maintaining the turnover of muscle proteins has been demonstrated by a point mutation in the
human kinase domain that causes myopathy (Lange et al., 2005). We hypothesized that the titin
kinase of smooth muscle performs the functions that have been proposed for striated muscle,
namely signal transduction and stretch sensor to regulate protein expression in a strain-
dependent manner via substrates (Tskhovrebova and Trinick, 2003; Granzier and Labeit, 2004;
Lange et al., 2005). Here, we show the presence of a titin transcript that includes Z-disc, N2A,
and M-band exons in uterine tissue (Figure 17). Other studies have confirmed the expression of
a titin transcript encoding the N-terminus in uterus by RT-PCR (Sorimachi et al., 1997; Labeit et
al., 2006; Chi et al., 2008). The transcription level of the M-band exons 1, 2, and 6 was similar
to the expression of the Z-disc exon 3 and the N2A exon and there was no transcript of titin’s M-
band exons 1 and 2 in our knockout mice. This led us to the conclusion that titin’s kinase region
is expressed in uterine tissue although results obtained by Labeit and coworkers suggest that
smooth muscle titin consists of Z-disc, I-band, and A-band parts with an alternative C-terminus
devoid of an M-band segment (Labeit et al., 2006). In this study human aorta, bladder, carotid,
and stomach have been investigated, but not uterus. Compared to striated muscle, the expression
level of smooth muscle titin is ~100 fold less (Labeit et al., 2006), which we also observed at the
Discussion
83
high cycle threshold value during our Real-Time PCR analysis. It is predicted that the extensible
I-band regions are short in smooth muscle titin (Labeit et al., 2006) so that stretch will result in
relatively high forces, which would compensate for smooth muscle titin’s low abundance.
To study the function of smooth muscle titin (Figure 15), we used a cre-recombinase under the
control of the smooth muscle myosin heavy chain promoter after demonstrating that it led to
recombination in uterine smooth muscle cells (Figure 16). We were able to efficiently delete
titin’s M-band exons 1 and 2 in uterus of knockout females in comparison to control mice
(Figure 17). Our knockout and control mice carried one universally recombined titin allele due
to germline activity of the cre-recombinase (Figure 17A). This is consistent with recently
published work, which shows that the cre-recombinase activity in male germ cells is
independent of the cre-genotype and ends prior to fertilization (de Lange et al., 2008). Hence
recombination occurred in all haploid male germ cells, in which the floxed allele was present,
but did not affect the female allele. As a result, the control and knockout animals of the
SMcTiMEx1-2 strain carried universally one deleted titin allele, whereas the smooth muscle
activity of the cre-recombinase caused also recombination of the second titin allele in the
knockout animals resulting in the complete loss of titin’s M-band exons 1 and 2. Heterozygous
mice contained one deleted allele in smooth muscle caused by the smooth muscle cre-activity.
Although heterozygously recombined (Figure 17B), the control and heterozygous mice still
expressed titin’s M-band exons 1 and 2 and did not display impaired implantation (Figure 19,
Figure 20, Figure 21, Figure 22). We also controlled that the cre-recombinase did not cause the
phenotype (Figure 22) as it has been reported for mammalian cells (Loonstra et al., 2001).
The smooth muscle specific loss of titin’s kinase region did not lead to an obvious phenotype
since the mice were viable and also had the same body weight as well as heart weight to
body weight ratio as their litter mate controls (Figure 21B, supplement Figure 31). An altered
heart weight to body weight ratio could indicate vascular defects that change the load on the
heart by altering the blood flow or that lead to dysfunction of the coronary vasculature (Bellomo
et al., 2000). The expression of the SMMHC cre-recombinase transgene starts in late embryonic
development at E12.5 (Xin et al., 2002). In addition, titin has a long half life of ~3 days (Isaacs
et al., 1989) so that the complete replacement of the wild-type with the mutant titin takes time.
Mice, in which the striated muscle specific deletion of titin’s kinase region started in late
embryonic development, did also not show an obvious phenotype until three weeks of age
(Gotthardt et al., 2003). At this time only 30-40% of total titin lacks the kinase region
(Ottenheijm et al., 2009a). Hence, a strong smooth muscle phenotype was not expected.
Moreover angiogenesis only contributes to organ growth after birth but most blood vessels
Discussion
84
remain quiescent during adulthood (Carmeliet, 2005). Then angiogenesis occurs only in the
female reproductive system and in response to pathological stimuli such as inflammation and
wound healing.
More understanding of preimplantation and implantation biology will help to improve fetal
health and female fertility. The SMcTiMEx1-2 strain was a suitable mouse model to challenge
the uterine tissue by pregnancy that involves smooth muscle contraction, angiogenesis, and
tissue remodeling. Our studies demonstrate a critical role in embryo implantation for titin’s
kinase domain and its surrounding regions, which both contain binding sites for multiple
proteins. The deletion of titin’s M-band exons 1 and 2 resulted in altered embryo spacing (Figure
19). Thereby the embryo implantation sites were mostly localized in only one uterine horn
(~24%) or crowded (13.6%). Furthermore, we observed 4 or fewer implantation sites in 22.7%
of knockout females. Additionally, the amount of implantation sites was significantly reduced
(Figure 20A). The aberrant implantation was confirmed for both E4.5 and E5.5, which also
verified on-time implantation in knockout females before E4.5. These data indicate that the loss
of titin’s kinase region could involve direct effects on the female reproductive system. To
explore this possibility, we investigated estrous cycle length, embryo transport through the
oviduct, and blastocyst development as major events in female reproduction (Figure 18). There
was no difference resulting in the same amount of E3.5 blastocysts available for implantation in
control and knockout animals. Furthermore, knockout uteri did not display hypoplasia as the
uterus weight to body weight ratio was comparable to the control group at E3.5 and E4.5 (Figure
21). Therefore the reduced number of implantation sites in knockout females was caused by an
impaired implantation. The amount of E3.5 blastocysts (Figure 18B) were determined by
flushing the uterine horns and counting. The average number of implantation sites (Figure 20A)
were determined by Evans Blue staining so that the number of both experiments differ.
The implantation phenotype upon loss of titin’s kinase region resembles the targeted deletion of
the genes encoding the LPA receptor LPA3 as well as the cytosolic phospholipase A2 (cPLA2)
(Ye et al., 2005; Song et al., 2002). A similar phenotype has also been reported for mice and rats
that were treated with indomethacin, which is an inhibitor of cyclooxygenases (Kennedy, 1977;
Kinoshita et al., 1985). These mouse models had a reduced litter size and altered embryo
spacing, but in contrast to our titin’s kinase region deficient mouse model also delayed
implantation. Additionally, cPLA2 knockout mice exhibited impaired preimplantational
embryonic development (Table 10). COX2 deficient mice have implantation defects as well, but
more severe other reproductive failures in ovulation, fertilization and decidualization (Lim et al.,
1997). In these previously described mouse models the prostaglandin biosynthesis was impaired,
Discussion
85
since cPLA2 is an important enzyme for the arachidonic acid production that is converted by
COX2 to the prostaglandin H2. Also the loss of LPA3 could be correlated to the cPLA2–
arachidonic acid–COX2–prostaglandin signaling pathway because reduced Cox2 expression and
prostaglandin levels in E3.5 pregnant uteri of this mouse model has been shown (Ye et al.,
2005). The deletion of titin’s kinase region did not alter the expression of Cox2 and Lpar3 in
E3.5 pregnant uteri compared to control females (Figure 27). Consistent with our findings, it has
not been possible to correct uneven embryo spacing and the decrease of embryo implantation
sites in cPLA2COX2, and LPA3 deficient mice by rescuing the prostaglandin reduction with
prostaglandin E2 and carbaprostacyclin administration (Song et al., 2002; Lim et al., 1999; Ye et
al., 2005). This rescue only restored on-time implantation indicating that prostaglandin
biosynthesis is crucial to determine the window of implantation and independent effects of
arachidonic acid are involved in embryo spacing and the implantation reaction, which is
impaired by the loss of titin’s kinase region.
Table 10: Knockout mouse models with an implantation phenotype.
Knockout model Reduced number
of implantation
sites
Mislocalization
and crowding
Delayed
implantation
Preimplantational
effects
titin’s kinase region yes yes no no
LPA3 yes yes yes no
cPLA2 yes yes yes yes (mild)
COX2 yes (severe) not detectable yes yes (severe)
L-12/15-LOX yes unknown no unknown
Another pathway that regulates implantation through arachidonic acid metabolism is described
in mice carrying a null mutation for gene encoding the L-12/15-LOX that catalysis the formation
of hydroxyeicosatetraenoic acids from arachidonic acid (Li et al., 2004). It has been shown that
the expression of Alox15 in uterus is induced by progesterone with a maximum before
implantation, which results in an increased uterine level of 12-HETE (Li et al., 2004). Therefore
Real-Time PCR, metabolite measurement, and proteomics were conducted with tissue from E3.5
pregnant mice. This time point also minimized changes due to secondary effects. Li and
coworkers have shown that deletion of the L-12/15-LOX caused a reduced number of
implantation sites at on-time implantation (Li et al., 2004). For this mouse model nothing is
reported about the localization of the implantation sites within both uterine horns (Table 10).
Discussion
86
Nevertheless, this implantation phenotype is markedly similar to the females lacking titin’s
kinase region (Figure 20). Consistently, both knockout models had reduced uterine levels of 12-
HETE before implantation (Figure 23B; Li et al., 2004). It has been shown that additional
inhibition of the epidermal 12/15-lipoxygenase (E-12/15-LOX) activity reduced the uterine 12-
HETE level and blocked embryo implantation nearly completely in L-12/15-LOX knockout
mice (Li et al., 2004). Hence the partial but not total loss of implantation sites can be explained
by the compensatory effect of the E-12/15-LOX that caused residual 12-HETE production in
uterine tissue. Li and colleagues also have uncovered that the reduction of embryo implantation
caused by an E- and L-12/15-LOX inhibitor could be reversed by a PPARagonist. These results
suggest that PPARis a downstream target of 12-HETE and important during early pregnancy.
We found in our knockout model of titin’s kinase region no upregulation of Pparg expression
(Figure 23A). Nevertheless, there might be a change on protein level or activation of the PPAR.
Hence, we will try to rescue the reduced number of implantation sites using a PPAR selective
agonist to ascertain if titin’s kinase region is related to this signaling pathway.
Although L-12/15-LOX and its metabolite 12-HETE have been identified as important regulator
of implantation, the precise biological events that caused the reduced number of embryo
implantation sites remain elusive. It needs to be considered that the L-12/15-LOX has a wide
distribution. It is not only expressed in the surface and glandular epithelial cells of the uterus,
but also in vascular smooth muscle cells (Li et al., 2004; Natarajan et al., 1993). 12-HETE has
been implicated in inflammation-related and other physiological pathways such as angiogenesis
that is completed by the recruitment of vascular smooth muscle cells (Connolly and Rose, 1998;
Nie et al., 2006). Angiogenesis is an essential physiological component of pregnancy because a
richly vascularized uterine tissue is needed for implantation and development of the placental
vasculature to facilitate the transport of nutrients and oxygen to the embryo. It occurs together
with increased vascular permeability at the site of blastocyst apposition and is extended during
the penetration phase (Plaks et al., 2006; Schlafke and Enders, 1975). The reduced number of
implantation sites in mice lacking titin’s kinase region is related to this phase because we could
not flush blastocysts from E4.5 knockout uteri indicating that apposition and attachment of
blastocyst occurred normally. Furthermore, we detected in one knockout uterus implantation
sites that have not further developed (Figure 20B) and the number of implantation sites in
knockout females was already reduced at E4.5 (Figure 20A). The knockout uteri also had a
delayed development during early pregnancy at E5.5 as measured by the uterus weight to body
weight ratio (Figure 21). This indicates that the deletion of titin’s kinase regions affected uterine
vascularization or decidualization. Consistent with these findings, the in vivo angiogenesis
Discussion
87
studies revealed decreased angiogenesis in response to VEGF in knockout mice (Figure 24A).
Thus, the reduced number of implantation sites appears to be at least in part due to impaired
angiogenesis, which is likely independent of VEGF as our knockout and control animals had the
same VEGF serum levels (Figure 24B). This is in agreement with results obtained by
Matsumoto and coworkers, who showed that VEGF directs uterine vascular permeability and
angiogenesis during the preimplantation and attachment phase driven by prostaglandins
(Matsumoto et al., 2002). Accordingly mice that carry a null mutation for the gene encoding
COX2 already exhibited implantation defects during the initial attachment phase (Lim et al.,
1997).
Our investigation revealed PDGF-B as growth factor that might be involved in vascular growth
and remodeling during early stages of pregnancy dependent on titin’s kinase region since the
PDGF-B serum levels were increased in pregnant knockout mice (Figure 24B). TNF-, another
angiogenic agent that is regulated at the time of implantation (De et al., 1993), was unchanged
indicating a specific role for PDGF-B. It is known that PDGF-B activates intracellular
phospholipases producing arachidonic acid, which is metabolized by lipoxygenases,
cyclooxygenases, and cytochrome P-450 oxygenases (Smith, 1989). It has been demonstrated
that PDGF-B induces L-12/15-LOX activity and Alox15 expression in vascular smooth muscle
cells (Natarajan et al., 1996). In addition, the L-12/15-LOX product 12-HETE causes
extracellular matrix production as well as vascular smooth muscle cell migration (Nakao et al.,
1982; Natarajan et al., 1994). Hence we tested if smooth muscle cell migration was altered. We
discovered that smooth muscle cells lacking titin’s kinase region did not migrate in response to
PDGF-B (Figure 25). These data together with the decreased uterine levels of 12-HETE indicate
that titin’s kinase region affected PDGF-B and its mediator 12-HETE and thereby cell migration.
This is consisting with previous findings that PDGF-induced chemotactic effects were
significantly blocked by lipoxygenase inhibitors (Natarajan et al., 1996; Gu et al., 2001; Patricia
et al., 2001). PDGF-induced migration was also reduced in vascular smooth muscle cells
obtained from the L-12/15-LOX knockout model (Reddy et al., 2003). Since angiogenesis
requires vascular smooth muscle cells, which are recruited to the site of vessel formation by
PDGF-B (Lindahl et al., 1997; Hellström et al., 1999), it is tempting to speculate that the
reduced number of implantation sites in titin’s kinase region and L-12/15-LOX knockout mice
was caused by impaired vessel formation or remodeling. The newly formed blood vessels were
not stabilized properly by the recruitment of smooth muscle cells. Therefore, embryos
implantation was hampered but could be partially rescued by the activity of the E-12/15-LOX
activity. The PDGF-B levels were increased to compensate the smooth muscle migration failure.
Discussion
88
Prospective experiments will be done to confirm the link of 12-HETE to titin’s kinase region.
We will try to rescue the migration phenotype of knockout smooth muscle cells by
overexpressing L-12/15-LOX or by 12-HETE supplement. Moreover we will use other
chemoattractants than PDGF-B to investigate, if the migration defect is specific to 12-HETE–
PDGF-B signaling. It might be a general defect since stretch sensor is one proposed function of
titin’s kinase (Tskhovrebova and Trinick, 2003; Granzier and Labeit, 2004; Lange et al., 2005).
The titin molecule is a possible candidate to sense stretch and to convert it into a biochemical
signal because it contains besides the kinase domain also elastic spring elements as well as
multiple protein binding and phosphorylation sites.
Expression analysis of genes encoding M-band interaction partners (Figure 29), whose binding
sites have been deleted in titin’s kinase region deficient animals, revealed no alteration of Nbr1
and p62 that are expressed in smooth muscle and encode postulated kinase substrates (Figure
26).
Figure 29: Binding partners of titin’s kinase region. The binding proteins of titin’s kinase region are shown
(structural proteins: purple; adapter proteins: yellow; signaling proteins green). The protein domains, which are
encoded by titin’s M-band exons 1 and 2 that are deleted in our knockout model, are also depicted (Ig domains
(red), FN3 domain (white), unique sequences (blue), kinase domain (black)). The domains of titin’s kinase region of
the A-band (A169, A170) and of the M-band (M1-M7) are specified. Modified from Gotthardt et al., 2003.
A small reduction of Fhl2 transcripts was observed (Figure 26). The knockout models of titin’s
PEVK and N2B segment also had only a little or no change, respectively, in the Fhl2 expression
in heart, but a strong effect on protein level (Granzier et al., 2009; Radke et al., 2007). Hence,
future studies will show, if there is a change in protein amount of uterine tissue obtained from
titin’s kinase region deficient mice. FHL2 will be investigated because it participates in wound
healing by contributing to gene expression, cytoarchitecture, cell adhesion, signal transduction,
and migration. Wound healing and pregnancy are related because they share the need for
angiogensis and remodeling of the extracellular matrix. Therefore, we will conduct wound
healing studies in the future. Of note, it has been shown that impaired wound healing in FHL2
knockout mice is due to disturbed collagen metabolism (Kirfel et al., 2008). Titin’s kinase region
deficient knockout uteri pregnant at E3.5 had an increased amount of collagen type VI alpha 2
Discussion
89
(COL6A2; Figure 28), which is involved in the decidualization of embryo implantation
(Mulholland et al., 1992). Hence, we will test the decidual reaction of our knockout females in
vivo by artificial induction of decidualization. Moreover it has been shown that FHL2
translocates to the nucleus triggered by LPA and S1P (Müller et al., 2002). LPA and S1P are both
important for successful implantation (Ye et al., 2005; Mizugishi et al., 2007). In addition,
sphingolipid metabolism regulates uterine decidualization and blood vessel stability and S1P is
required for PDGF-B guided migration of vascular smooth muscle cells during angiogenesis
(Liu et al., 2000; Allende and Proia, 2002; Spiegel and Milstien, 2003).
Murf1 had an increased transcription level in pregnant uteri that were deficient in titin’s kinase
region (Figure 26). Further studies will provide insight if the M-band binding protein MuRF1 is
also upregulated on protein level. Despite the deletion of the MuRF1 binding site, a higher
MuRF1 protein amount has been detected in adult heart lacking titin’s kinase region (Peng et al.,
2007). The increased level of the ubiquitin ligase MuRF1 might result in increased
ubiquitination and thereby in alteration of heat shock response and protein degradation. Previous
studies suggest that the ubiquitin-proteasome pathway effects embryo implantation probably by
being involved in degradation and remodeling of the uterine tissue during decidualization (Wang
et al., 2004). Indeed, we observed upregulation of the DnaJ (Hsp40) homolog (DNAJA1) and
downregulation of the heat shock protein 1 (HSPB1), the protein disulfide-isomerase A4
(PDIA4), and the proteasome 26S subunit ATPase 2 (PSMC2; Figure 28) on protein level.
Components of the ubiquitin-proteasome pathway were also altered in striated muscle specific
titin’s kinase region knockout mice (Peng et al., 2006). The reduced amount of uterine 12-HETE
in our knockout mice that is contrary to the upregulated transcriptional level of Alox15 (Figure
23) might be caused by variation of the ubiquitin-dependent protein degradation via proteasomes
leading to a reduction in L-12/15-LOX protein amount. There could also be a link between
MuRF1 and FHL2 because it has been shown that the FHL2 protein level is increased in mice
deficient for both MuRF1 and MuRF2 (Witt et al., 2008).
Beside the effect on heat shock response and protein degradation, the proteins that were changed
in E3.5 pregnant knockout uteri mainly belong to metabolism and the cytoskeleton (Figure 28).
This is consisting with findings from quadriceps of the striated muscle specific titin’s kinase
knockout (Raddatz et al., 2008). Differential regulation of proteins of the cytoskeleton such as
transgelin (TAGLN), actin related protein 2/3 complex (ARPC2), capping protein (CAPG), and
vimentin (VIM) support the hypothesis that deletion of titin’s kinase region caused altered tissue
remodeling in early pregnancy as already indicated by an impaired ubiquitin-proteasome
pathway. Fibrinogen (FGA) and the splicing factor arginine/serine-rich 3 (SFRS3), which were
Discussion
90
accumulated in pregnant knockout uteri (Figure 28), have been related to implantation in
previous studies. It has been demonstrated that fibrinogen is crucial for maintenance of
pregnancy by supporting development of fetal-maternal vascular communication and by
stabilizing embryo implantation (Iwaki et al., 2002). The splicing factor arginine/serine-rich 3
was identified as marker for uterine receptivity (Reese et al., 2001). The findings confirm the
implantation defect in titin’s kinase knockout mice.
To gain more insight into the signaling pathways that are affected in our knockout mice, we
investigated cytokines in uterus and serum of pregnant mice. We focused on cytokines that have
been related to implantation in mice but also to recurrent miscarriage of women and a cytokine
profile that is predictive for in vitro fertilization (Jasper et al., 2007; Boomsma et al., 2009). The
tested cytokines were not altered (supplement Figure 32, Figure 33). Most remarkable, LIF that
is a key regulator for implantation (Stewart et al., 1992) was unchanged indicating that
implantation normally initiated in titin’s kinase knockout animals. This is similar to the LPA3
knockout mouse model that also exhibited impaired implantation (Ye et al., 2005).
Relevant to our work were also previous experiments that have been done with smooth muscle
cells obtained from L-12/15-LOX knockout mice. It has been shown that they had reduced
activation of the mitogen-activated protein kinase 14 (MAPK14) and the CREB1, which both
mediate cellular effects induced by 12-HETE (Reddy et al., 2002). MAPK14 knockout females
have defects in placental angiogenesis leading to embryonic death (Mudgett et al., 2000; Adams
et al., 2000). In contrast to the transcript level of Creb1 that was unchanged, we found in uteri
from our knockout animals increased expression of Mapk13 (Figure 23A). Mapk13 encodes the
MAPK13, which is homologous to MAPK14 but has different substrate specificity (Hu et al.,
1999). Future studies will expose if the activation of the kinases MAPK13 and MAPK14 occurs
differently upon loss of titin’s kinase region in pregnant uterus thereby modulating signaling
pathways that might be responsible for altered growth factor response via the mediator 12-
HETE.
So far, we investigated the pathway leading to a reduced number of implantation sites. The
molecular mechanisms underlying crowding and mislocalization of embryo implantation sites,
which are most likely related to smooth muscle contraction, are unknown. It has been shown that
titin’s kinase region acts a as regulator of contractile function by calcium handling (Peng et al.,
2007). The expression of selected genes encoding calcium related proteins (supplement Figure
34) did not differ in titin’s kinase region deficient pregnant knockout uteri. Also the transcript
level of Lpar3 encoding LPA3, which has been shown to mediate embryo spacing as well as
implantation via different signaling pathways (Hama et al., 2007), was unchanged (Figure 27). It
Discussion
91
might be that LPA3 is altered on protein level or that titin’s kinase region is downstream of
LPA3 signaling. Hence, additional studies are required to understand smooth muscle titin’s
mechanical and signaling function. Protein kinase G (PKG) will be investigated because titin
and the actin related protein 2/3 complex (ARPC2), which is upregulated on protein level in
pregnant knockout uteri (Figure 28), were identified to be in vitro PKG substrates in pregnant rat
uteri (Huang et al., 2007). Using our proteomics approach, we will also generate a protein
reference map for pregnant mouse uterus after analyzing more protein spots. Identification of
proteins based on their migration in 2D-gels will provide a molecular basis for functional and
pathophysiological studies that will help to elucidate pregnancy in more details.
To investigate titin’s kinase region deficient mice in pregnancy, timed matings were set up so
that the genotypes of the embryos, which developed in control and knockout females, were
similar. Hence, knockout males were bred with control females and control males were bred
with knockout females. However, activity of the cre-recombinase before and after fertilization in
knockout females caused recombination of both titin alleles independent of the genotype of the
embryo in 100% (de Lange et al., 2008). In control females, the cre-recombinase activity in the
male germ cells ended prior to fertilization. This led to ~50% of embryos with universal deletion
of both titin alleles taken into account that control and knockout animals carried one universally
recombined allele. Since 100% of the embryos that developed in knockout females and only
~50% of the embryos that developed in control females were deficient in titin’s kinase region,
blastocyst transfer experiments will be done to distinguish maternal from embryonic effects. The
lack of titin’s kinase region in the embryo itself might contribute to the phenotype because
results obtained from Du and colleagues indicate that the trophectoderm of the blastocyst, which
mediates the implantation of the embryo into the endometrium, expresses titin (Du et al., 2007).
Hence, titin has been implicated in being involved in promoting human trophoblast growth and
invasiveness (Du et al., 2007; Xie et al., 2010). Additionally, Xie and coworker could correlate
enhanced invasiveness of trophoblasts with titin expression and speculate that a lack in
invasiveness is related to miscarriage in human (Xie et al., 2010). Other results show altered
expression of titin in chorionic villous samples of embryos (Farina et al., 2009). These samples
were obtained from women, who have developed later in pregnancy preeclampsia (pregnancy-
induced hypertension), highlighting the importance of titin at the embryo-maternal interface.
These results suggest that the trophoblast of the blastocyst, which is deficient in titin’s kinase
region, could contribute to the reduction of embryo implantation sites.
Discussion
92
Our data provide the first genetic evidence for a role of smooth muscle titin because embryo
implantation studies indentified changes in titin kinase deficient females in comparison to
control mice. The aberrant implantation demonstrated at E4.5 and E5.5 resulted in
mislocalization and crowding as well as a reduced number of embryo implantation sites. We
related the decreased number of implantation sites to the penetration phase because the estrous
cycle length, embryo transport through the oviduct, blastocyst development, and blastocyst
attachment occurred normally. Loss of titin’s kinase region caused altered expression of the
genes encoding the M-band binding proteins MuRF1 and FHL2 as well as the L-12/15-LOX.
Consistently, we found an influence on arachidonic acid metabolism with a decreased level of its
metabolite 12-HETE in pregnant uterus. Angiogenesis was reduced in knockout mice, whereas
we identified PDGF-B as potential angiogenic factor that is altered by the loss of titin’s kinase
region. Knockout smooth muscle cells did not migrate in response to PDGF-B. Therefore we
speculate that the impaired angiogenesis was due to a failure in stabilizing nascent vessels
because the recruitment of smooth muscle cells to the site of vessel formation was hampered.
Since angiogenesis is required for embryo implantation, aberrations in these processes might
contribute to pregnancy loss. Hence it is tempting to speculate that the angiogenesis defect in
our knockout mice is the reason for the reduced number of implantation sites because some
embryos were not able to implant properly. This is the first link of the titin’s kinase region to
PDGF-B and its mediator 12-HETE. Thereby, titin was identified as a new molecular regulator
of angiogenesis during early pregnancy, which influences female fertility. So far, impaired
vascular development that may lead to implantation failure and early miscarriage in humans has
not been well studied, although disruption of angiogenesis at the site of implantation is
associated with poor reproduction in humans (Meegdes et al., 1988; Vuorela et al., 2000).
Moreover, it is also possible that complications in later pregnancy, which affect fetal health
(intrauterine growth restriction) and maternal health (preeclampsia), are associated with
molecular and cellular implantation defects (Reynolds et al., 2006). Hence, a better
understanding of implantation biology will improve female fertility and fetal health because it
will allow novel therapeutic advances. Tissue availability restricts studies in human, but mouse
models are a suitable tool of embryo implantation because similarities between species exist.
Using the knockout model of titin’s kinase region, we discovered a potential molecular
mechanism of angiogenesis during early pregnancy in vivo, which needs to be carefully
interpreted in addressing human fertility and clinical relevance.
Outlook
93
7 Outlook
The DsRed knockin mouse model that we generated will not only help to visualize smooth
muscle titin, but also to study titin in striated muscle. Our objective is to uncover the molecular
mechanisms underlying sarcomere assembly, disassembly, and turnover in myofibrillogenesis
and adulthood, which is important in the physiological adaptation of muscle growth and in
disease. Because of titin’s large size, it is not possible to express titin tagged with a fluorescence
protein in cell culture as it has already been done for several sarcomeric proteins (Wang et al.,
2005). Hence our mouse model provides a novel approach to visualize titin's sequential
integration into and disintegration from the sarcomere at different stages of development in vivo
using live cell imaging. The analysis of protein turnover as well as mobility of titin along the
sarcomere in vivo will be done utilizing photobleaching.
The smooth muscle specific knockout of titin’s kinase region displayed an implantation
phenotype. There is still a need to unravel the pathways responsible for smooth muscle
contraction that led to crowding and mislocalization of embryo implantation sites. Fetal and
maternal health, e.g. by reducing ectopic pregnancies, placenta previa, as well as pre-term labor,
will benefit from an increased knowledge of uterine contraction at cellular and molecular level.
So far, we analyzed the molecular mechanism underlying the reduced number of implantation
sites indicating that titin’s kinase region is involved in angiogenesis via PDGF-B. Angiogenesis
in adults is crucial for embryo implantation and wound healing, but an imbalance in
angiogenesis also contributes to malignant, inflammatory, ischemic, infectious, and immune
diseases. Hence, the increased understanding of angiogenesis will help to improve fetal health
and female fertility as well as therapeutic strategies, such as revascularization of ischemic
tissues or inhibition of angiogenesis in cancer and skin disorders. The relevance of PDGF-B
signaling as a therapeutic target has been demonstrated because inhibition of the PDGF-B and
VEGF pathway in a transgenic mouse model of pancreatic islet tumors resulted in
antiangiogenic and antitumor effects (Bergers et al., 2003). Furthermore, it has been shown that
the inhibition of PDGF-B signaling reduced interstitial tumor pressure thereby enhancing the
effect of chemotherapy (Pietras et al., 2001, 2002). We will investigate the role of titin in
angiogensis in more detail focusing on smooth muscle migration in response to PDGF-B, which
might be important for angiogenesis in health and disease.
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9 Supplement
Figure 30: Map of the targeting vector DsRedKI. The targeting vector had a size of 15347 bp spanning the
genomic region from titin’s exon 18 to exon 31 with the DsRed incorporated in exon 28 and the neomycin
resistance cassette with the glucokinase promoter (GK) flanked by 2 FRT-sites between exon 28 and exon 29. The
plasmid also includes the pUC origin for propagation in E.coli, the T7 promoter, and the ampicillin resistance gene
(ampr) for antibiotic selection in E.coli. The restriction site of ScaI that linearizes the targeting vector is indicated.
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Figure 31: Adult smooth muscle knockout animals did not have an obvious phenotype. A) Deletion of titin’s
kinase region in knockout (KO) animals did not change the heart weight to body weight ratio in comparison to
control (CON) animals, which was shown for E3.5 (KO n=10; CON n=11), E4.5 (KO n=5; CON n=7), and E5.5
(KO n=7; CON n=6; mean with SEM; Two-way ANOVA P>0.05). E3.5 was confirmed by the presence of
blastocysts. B) Control, heterozygous (HET), knockout, and Cre+ control mice (Cre+) had the same body weight so
that the reduced uterine weight to body weight ratio in knockout animals was due the loss of titin’s kinase region
(CON n=6; HET n=5; KO n=7; Cre+ n=3; mean with SEM; one-way ANOVA P>0.05).
Figure 32: Increase in the PDGF-B serum levels was not accompanied by changes in selected cytokines.
Multiplex bead immunoassay to measure cytokines in pregnant E3.5 mice revealed no difference in the serum levels
of G-CSF, IL-6, IP-10, and TNF- between knockout animals (KO; n= 13) and control animals (CON; n=14; mean
with SEM; T-test P>0.05). The metabolite concentration is illustrated per serum volume.
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Figure 33: LIF, an important factor of embryo implantation, was unchanged in pregnant E3.5 knockout
uteri. The cytokines IL-1, IL-6, IP-10, LIF, MCP-1, and TNF- were unchanged in pregnant E3.5 uteri comparing
knockout (KO; n=13) and control animals (CON; n=14; mean with SEM; T-test P>0.05). Values were determined
using the multiplex bead immunoassay. The metabolite concentration is illustrated per lysate volume that was
normalized against its total protein concentration.
Figure 34: The transcript level of genes encoding selected calcium-related proteins was unchanged. Real-Time
PCR analysis revealed that the expression of the genes Prkcd, Prkce, S100a1, Cacna1c, Camk2a, and Calb1
encoding selected calcium-related proteins did not differ between knockout (KO; n=3) and control (CON; n=3) uteri
pregnant at E3.5. The mRNA levels were normalized to 18S RNA and expression is displayed as fold change
relative to the control (mean with SEM; T-test comparing CON and KO of one primer/probe set P>0.05).
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Figure 35: Dual view of knockout and control 2D-gel. 2D-gel electrophoresis was performed for proteome
analysis of pregnant E3.5 knockout (KO; n=6) in comparison to control (CON; n=6) uteri. A dual view of a
representative control and knockout gel is shown, which was generated using the Delta2D software. Thereby, the
software colorized the control protein spots green and the knockout protein spots red leading by overlay in the dual
view to yellow spots, if the same amount of protein is present in both groups. Proteins in knockout samples that are
downregulated are indicated by green spots (32 proteins) and that are accumulated (74 proteins) are indicated by red
spots in the dual view (T-test P<0.1 *).
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Table 11: Differentially regulated proteins in E3.5 pregnant knockout uteri. The table shows the mean of
control (CON n=6) and knockout (KO n=6) percent volume value (% V) of protein spots obtained by evaluation of
2D-gels comparing pregnant E3.5 knockout and control uteri. Furthermore, the %V ratios between knockout and
control mean as well as the results of the T-test are listed. Thereby, proteins with a ratio value of < 0.8 and > 1.3
were considered to be less or more synthesized, respectively (T-test P<0.1 *).
Protein Protein name
Mean
% V
CON
Mean
% V
KO
Ratio
% V
KO/
CON
T-test
ANXA2 annexin A2 0.108 0.084 0.78 0.006
ARPC2 actin related protein 2/3 complex 0.081 0.11 1.36 0.037
CAPG capping protein 0.029 0.042 1.43 0.064
CAR2 carbonic anhydrase 2 0.068 0.091 1.34 0.011
CKB brain creatine kinase 0.233 0.18 0.77 0.01
COL6A2 collagen type VI alpha 2 0.069 0.101 1.46 0.074
COL6A2 collagen type VI alpha 2 0.042 0.076 1.79 0.009
DNAJA1 DnaJ (Hsp40) homolog 0.03 0.045 1.49 0.038
ETFDH electron transferring flavoprotein dehydrogenase 0.031 0.043 1.37 0.024
FGA fibrinogen 0.032 0.05 1.57 0.014
HNRNPAB heterogeneous nuclear ribonucleoprotein A/B 0.034 0.05 1.48 0.036
HSPB1 heat shock protein 1 0.191 0.141 0.74 0.024
NDUFS3 NADH dehydrogenase (ubiquinone) Fe-S protein 3 0.03 0.068 2.24 0.003
PCX pyruvate carboxylase 0.029 0.044 1.57 0.026
PDIA4 protein disulfide-isomerase A4 0.253 0.205 0.79 0.015
PSMC2 proteasome 26S subunit ATPase 2 0.039 0.026 0.68 0.019
SFRS3 splicing factor arginine/serine-rich 3 0.05 0.068 1.35 0.029
TAGLN transgelin 0.498 0.4 0.79 0.016
TPI1 triosephosphate isomerase 1 0.167 0.217 1.3 0.021
VIM vimentin 0.018 0.027 1.48 0.05
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