Formation of lipid nanodiscs through SMALPs to extract membrane proteins in their native lipid environment

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To all my family:

To my father, for being a super father, for helping me in any possible way, for never leaving me behind, for all the support, for all the patience to conversations of hours and hours, for doing everything in his hands for me, for loving me unconditionally and for being the most close to perfection that I know.

To my brother who I always say that is annoying, but I love him anyway. For growing with me and even for annoying me so much, that raised my patience levels to something I never imagined.

To my mother, for being the person who helped me the most shaping the person I am today and for teaching me everything I know about every little thing.

To my aunt Fernanda for being such a well-humoured person, for always making me laugh and all the conversations. To my uncle Rui and my cousin Emmanuel for being as jokers as I am, for following my silly jokes and making me laugh until I can’t stand the pain in my cheeks and abdominals.

To my grandfather Aires who I love to discuss philosophy with, for being such a kind hearted person, for hearing my outbursts and for always receiving me with a huge smile and with arms wide open to hug me.

To my other grandparents Laurinda, Miquelina and Garrido for raising me while I was a child, for always telling me stories of their lives when I was a little and rebel kid, that always made them laugh.

To my cousin Ângela, for supporting me unconditionally every time I needed. For being such a great support in such a difficult time in my life and always calling me “family genius”.

To my supervisor Prof. Margarida Bastos, for receiving me again in her lab team, for the advices at the right time, for the patience and availability to help me and teach me new things again.

To my lab team, Bárbara Claro, Inês Martins, Regina Adão and all the other people present in the physical chemistry floor, for always helping me when I needed.

To my co-supervisor, Prof. Maria João Moreno and Patrícia Martins, from Faculdade de Ciências e Tecnologia de Coimbra, for the opportunity to collaborate and work in another lab environment, for all the help and patience along my work while in

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To all my close friends:

To Rita Magalhães, my “forever lab partner” which has been doing this path with me since we were biochemistry students, for all the time she was present to help me and advise me.

To Laura, Joana and Raquel, my friends who supported me in the last months when I needed the most, that showed available for helping me at any time and for having a great and enormous heart.

To Adelaide Sousa, Ana Pereira, Catarina Pelicano, Daniel Sousa, Inês Lorga, Inês Rocha, Joana Ferreira, José Pinto, Olena Shevchuk, Patrícia Soares, Pedro Lopes, Pedro Nicola, Ricardo Esteves, Sara Vila Maior and Sara Fialho for all the patience to listen to me when I needed the most, and all the support. For always providing me with a great time with them, all the healthy laughs, all the kindness and for always being there for me.

To my break and coffee partners Dmitriy and Clara that always made me laugh when I needed the most and showed me support every time.

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Nas últimas décadas verificou-se uma maior investigação na área da proteómica.

Para estudar proteínas de membrana é muitas vezes necessário extraí-las de modo a que se possam realizar experiências em condições in vitro controladas. Tradicionalmente, a extração de proteínas membranares é feita através de detergentes. Este processo é eficiente, embora, por vezes, leve à desnaturação das proteínas e/ou perda de função, impossibilitando assim o seu estudo posterior. Mas a principal deficiência do processo de extração com detergentes é o facto de a proteína deixar de estar no seu ambiente lipídico natural.

A alternativa apresentada neste trabalho, para uma extração eficaz, de proteínas membranares, mantendo o seu ambiente lipídico natural, envolve o uso de co-polímeros que formam nanodiscos lipídicos. Este processo permite extrair as proteínas no seu ambiente lipídico nativo. O uso de co-polímeros para extração de proteínas membranares é recente, mas tem-se mostrado bastante promissor. No presente trabalho este processo foi usado para a extração de uma glicoproteína-P (P-gp), da membrana do inseto da espécie Spodoptera Frugiperda (Sf9).

Numa fase inicial foram realizadas experiências metodológicas, usando o co-polímero em membranas modelo (lipossomas), de forma a optimizar as condições experimentais e ganhar conhecimento do sistema e processos envolvidos. Nestas experiências foram testados os co-polímeros de estireno (S) – ácido maleico (MA) SMA 2:1 e SMA 3:1, em sistemas modelo de DPPC:DMPS (7:3), DPPC:DMPS (6:4), DPPC:DMPG (3:1), POPE:POPG (7:3) e POPC:POPE:Cardiolipina (55:35:10). Numa segunda fase, tentou-se fazer a extração da P-gp no seu ambiente nativo na membrana da espécie Spodoptera Frugiperda, usando o co-polímero SMA 3:1. A formação de nanodiscos em membranas modelo foi seguida por DLS e DSC. No caso da extração da P-gp foi também usado o DLS para seguir a evolução de tamanho e foram realizados ensaios para avaliar a atividade da P-gp após extração (ensaios de ATPase – atividade proteica) e foi quantificada a concentração de proteína extraída (método de quantificação proteica).

Palavras-chave: proteínas, extração, co-polímero, nanodiscos, Spodoptera Frugiperda, P-glicoproteína, lipossomas, SMA

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In the last decades research has significantly increased in the area of proteomics.

To study membrane proteins it is often necessary to extract them in order to perform the needed experiments and assays in a controlled in vitro environment. Traditionally, the extraction of membrane proteins is made through detergents. This method is efficient, however leads, sometimes, to proteins’ denaturation and/or loss of function, making it impossible to do further studies. Nevertheless the main flaw of the extraction method with detergents is the fact that the protein loses its natural lipid environment.

The alternative presented in this work allows an efficient extraction of membrane proteins involving the use of copolymers that extract the membrane proteins into lipid nanodiscs, keeping thus their native lipid environment. The use of copolymers for protein extraction is recent, but has already shown to be very promising. In the present work, this process was used for the extraction of P-glycoprotein (P-gp), from the membrane of the insect of the species Spodoptera Frugiperda (Sf9).

Prior to P-gp extraction methodological experiments were made using the copolymer in model membranes (liposomes) in order to optimize the experimental conditions and gain knowledge about the system and the processes involved. In these experiments the copolymers styrene (S) – maleic acid (MA) SMA 2:1 and SMA 3:1 were tested, in model membranes of DPPC:DMPS (7:3), DPPC:DMPS (6:4), DPPC:DMPG (3:1), POPE:POPG (7:3) and POPC:POPE:Cardilipin (55:35:10). After this preliminary work, the extraction of P-gp was tried from the membrane of

Spodoptera Frugiperda, using the copolymer SMA 3:1. The nanodiscs formation in

model membranes was followed by DLS and DSC, and in the case of the extraction of P-gp DLS was also used to follow the size of the species formed and the protein activity (ATPase assays) after extraction was assessed and the concentration of extracted protein (protein quantification method).

Key words: proteins, extraction, copolymer, nanodiscs, Spodoptera Frugiperda,

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List of tables ...xv

List of abbreviations ... xvii

1. Introduction ... 18

1.1. The importance of cell membranes and membrane proteins ... 18

1.2. Cell membranes’ composition ... 18

1.3. Introduction to protein extraction ... 19

1.4. Protein extraction methods ... 20

1.4.1. Non-bilayer systems ... 22

1.4.1.1. Detergents ... 22

1.4.1.2. Amphipols ... 23

1.4.2. Bilayer systems ... 24

1.4.2.1. Bicelles ... 24

1.4.2.2. Nanodiscs bounded by membrane scaffold proteins ... 25

1.4.2.3. Nanodiscs stabilized by copolymers ... 26

1.5. P-glycoprotein ... 28

1.6. Main goal of present work ... 30

2. Experimental procedure ... 31

2.1. Buffers’ preparation ... 31

2.2. Copolymers’ preparation... 32

2.3. Preparation of model membranes ... 33

2.4. Lipid quantification ... 35

2.5. Preparation of lipid nanodiscs ... 35

2.6. Crude membranes of Spodoptera Frugiperda (Sf9) ... 37

2.6.1. Experiment I – testing the influence of time and pH ... 37

2.6.2. Experiment II – testing the influence of a different buffer ... 38

2.6.3. Experiment III – comparing the extraction ability of the copolymer and of the detergent DDM ... 39

2.6.4. Experiment IV – testing the influence of different temperatures ... 39

2.6.5. Experiment V –repeating experiment IV only at 4ºC and 20ºC ... 40

2.7. Protein quantification method ... 40

2.7.1. Protein quantification of BSA and CM – testing the interference of SMA 3:1 in the protein quantification method ... 41

2.7.2. Protein quantification of the CMs and the protein extracted with the detergent DDM ... 42

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2.9.1.1. Principles and used procedure ... 44

2.9.2. Differential Scanning Calorimetry (DSC) ... 45

2.9.2.1. Principles ... 45

2.9.2.2. Procedure ... 46

3. Results and discussion ... 48

3.1. Model membranes ... 48

3.1.1. DLS results and discussion ... 48

DPPC:DMPS (7:3) and SMA 3:1 ... 48

DPPC:DMPG (3:1) and SMA 3:1 ... 51

POPC:POPE:Cardiolipin (55:35:10) and SMA 3:1 ... 52

Overall comments ... 53

3.1.2. DSC results and discussion ... 54

DPPC:DMPG (3:1) and SMA 3:1 ... 57

3.1.3. DLS and DSC results for POPE:POPG (7:3) and SMA 3:1 ... 58

3.2. Protein extraction from native membranes of Spodoptera Frugiperda (Sf9) ... 62

3.2.1. Results of the interference of SMA 3:1 with the protein quantification method ... 62

3.2.2. Results of the experiment I – testing the influence of time and pH ... 63

3.2.3. Results of the experiment II – testing the influence of a different buffer ... 70

3.2.3.1. Results of the DLS measurements – experiment II ... 70

3.2.3.2. Results of the protein quantification – experiment II ... 72

3.2.4. Results of the experiment III – comparing the extraction ability of the copolymer and of the detergent DDM ... 73

3.2.4.1. Results of the DLS measurements – experiment III ... 73

3.2.4.2. Results of the protein quantification – experiment III ... 76

3.2.5. Results of the experiment IV – testing the influence of different temperatures ... 77

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3.2.6.2. Results of protein quantification – experiment IV and V ... 95

3.2.6.3. ATPase assays’ results – experiment V ... 96

3.2.6.3.1. Experiments at 4ºC ... 97

Control sample – pellet ... 97

Control sample – supernatant ... 99

DDM sample – pellet ... 101

DDM sample – supernatant ... 101

SMA sample – pellet ... 103

SMA sample – supernatant ... 104

3.2.6.3.2. Experiments at 20ºC – only the supernatant ... 104

DDM sample ... 104

SMA sample ... 106

4. Conclusion ... 107

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Figure 1.2. – Representation of porin’s crystals. Figure from [8] ... 20 Figure 1.3. – Representation of some extraction systems used for proteins, adapted

from [6]. Being a – surfactants, b – amphipols, c – bicell, d – membrane scaffold proteins and e – copolymers ... 21

Figure 1.4. – Molecular representation of dodecyl maltoside ... 23 Figure 1.5. – Molecular representation of sodium dodecyl sulphate ... 23 Figure 1.6. – Molecular representation of 3-[(3-cholamidopropyl)

dimethylammonio]-1-propanesulfonate ... 23

Figure 1.7. – Representation of (A) annular lipids arrangement (yellow) around a

protein representation (green) and (B) non-annular lipids (red) arranged nearby a representation of a protein (green). The surrounding lipids (blue) represent bulk lipids ... 24

Figure 1.8. – Representation of a vesicle, being the blue forms the lipids in their

constitution ... 25

Figure 1.9. - Representation of a bicell, being the blue spheres the lipids and the white

ones the detergents’ heads ... 25

Figure 1.10. – Molecular representation of styrene ... 26 Figure 1.11. – Molecular representation of maleic acid in the protonated form ... 26 Figure 1.12. – Reactional scheme of the synthesis of SMA copolymer, adapted from

[6] ... 26

Figure 1.13. – Representation of the sequence of the SMA copolymer ... 27 Figure 1.14. – Representation of the addition of SMA in a lipid suspension and the final

result. The copolymer is able to form nanodiscs that contain portions of the membrane, forming SMALPs. Figure adapted from [36]. ... 27

Figure 1.15. – Molecular representation of diisobutylene ... 28 Figure 1.16. – Molecular representation of diisobutylene/maleic acid ... 28 Figure 1.17. – Molecular representation of P-gp and its domains, as well as the

representation of the lipid membrane in grey lines. Adapted figure from [39] ... 30

Figure 2.1. – Molecular representation of the used lipids during experiments, a)

DMPG, b) DMPS, c) DPPC, d) POPC, e) POPE, f) POPG and g) cardiolipin ... 33

Figure 2.2. – Photography of the extruder used along the experimental work for the

extrusion of model membranes ... 34

Figure 2.3. – Comparison between liposome suspension (left) and liposome with

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Figure 2.4. – Representation of the disposition of the filters and the petri dishes. ... 42 Figure 2.5. – Representation of the filters after step 7 ... 42 Figure 2.6. – Simplified representation of a heat compensation calorimeter ... 46 Figure 3.1. – Representation of the size evolution of lipid nanodiscs of SMA 3:1 and

liposomes of DPPC:DMPS (7:3) ... 49

Figure 3.2. – Representation of the size evolution of lipid nanodiscs of SMA 2:1 and

liposomes of DPPC:DMPG (3:1) ... 52

POPC:POPE:Cardiolipin (55:35:10) ... 53

Figure 3.6. – Cp vs temperature for the up-scans for liposomes of DPPC:DMPS (7:3) (black) and SMA 3:1/liposomes mixture, with P/L ratios of 0.118 (red), 0.344 (green), 0.575 (dark blue), 1.15 (pink) and 2.29 (yellow) ... 54

Figure 3.7. – Cp vs temperature for the up-scans for liposomes of DPPC:DMPS (7:3) (black) and SMA 2:1/liposomes mixture, with P/L ratios of 0.116 (red), 0.345 (green), 0.573 (dark blue), 1.15 (pink) and 2.29 (yellow) ... 55

Figure 3.8. – Cp vs temperature for the up-scans of DPPC:DMPS (6:4) liposomes (black) and lipid nanodiscs of SMA 3:1/DPPC:DMPS (6:4) with P/L ratios of 0.100 (red), 0.300 (green), 0.500 (dark blue), 1.00 (pink) and 2.00 (yellow) ... 56

Figure 3.9. – Cp vs temperature for the up-scans for liposomes of DPPC:DMPG (3:1) (black) and SMA 3:1/liposomes mixture, with P/L ratios of 0.102 (red), 0.208 (green), 0.556 (dark blue), 1.25 (light blue) and 2.16 (pink) ... 57

liposomes of POPE:POPG (7:3) ... 59

Figure 3.11. – Cp vs temperature for the up-scans for liposomes of POPE:POPG (7:3) (black) and SMA 3:1/liposomes mixture, with P/L ratios of 0.1 (a), 0.15 (b), 0.2 (c), 0.3

(d) and 0.5 (e). First scan is in red, second scan is in green, third scan is in dark blue

and fourth scan is in pink ... 60

Figure 3.12. – Representation of the correlogram of the sample with SMA and the

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control, 48h after the addition of the copolymer, for experiment I ... 67

Figure 3.15. – Representation of the correlogram of the sample with SMA and the

control, 72h after the addition of the copolymer, for experiment I ... 69

Figure 3.16. – Representation of the correlogram of the sample with the control and

SMA, 2h after the addition of the copolymer, for experiment II ... 71

Figure 3.17. – Representation of the correlogram of the sample with the control and

SMA, after the centrifugation, for experiment II ... 72

Figure 3.18. – Representation of the correlogram of the sample with the control, DDM and SMA, 2h after the addition of the detergent and the copolymer, for the experiment III ... 74

Figure 3.19. – Representation of the correlogram of the sample with the control, DDM

and SMA, 24h after the addition of the detergent and the copolymer, for experiment III ... 75

Figure 3.20. – Representation of the correlogram of the sample with the control, DDM

and SMA, after the centrifugation, for experiment III ... 76

Figure 3.21. – Representation of the correlogram of the control samples, at 4ºC (blue),

20ºC (red) and 40ºC (green), at 2h, for experiment IV ... 78

Figure 3.22. – Representation of the correlogram of the effect of DDM in CM, at 4ºC

(blue), 20ºC (red) and 40ºC (green), at 2h, for experiment IV... 78

Figure 3.23. – Representation of the correlogram of the effect of SMA in CM, at 4ºC

(blue), 20ºC (red) and 40ºC (green), at 2h, for experiment IV... 79

Figure 3.24. – Representation of the correlogram of the control samples, at 4ºC (blue),

20ºC (red) and 40ºC (green) at 24h, for experiment IV... 80

(blue), 20ºC (red) and 40ºC (green), at 24h, for experiment IV ... 80

(blue) and 20ºC (red), at 24h, for experiment IV ... 81

Figure 3.27. – Representation of the correlogram of the control samples, at 4ºC (blue)

and 20ºC (red), and 40ºC (green) at 48h, for experiment IV ... 82

(blue), 20ºC (red) and 40ºC (green), at 48h, for experiment IV ... 83

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(blue), 20ºC (red) and 40ºC (green), after centrifugation, for experiment IV ... 85

(blue), 20ºC (red) and 40ºC (green), after centrifugation, for experiment IV ... 86

and 20ºC (red), 2h after the samples’ preparation, for experiment V ... 88

(blue) and 20ºC (red), 2h after the samples’ preparation, for experiment V ... 89

and 20ºC (red), 24h after the samples’ preparation, for experiment V ... 91

and 20ºC (red), after centrifugation, for experiment V ... 93

(blue) and 20ºC (red), after centrifugation, for experiment V ... 93

(blue) and 20ºC (red), after centrifugation, for experiment V ... 94

Figure 3.42. – Representation of the absorbance of the pellet of the control sample at

4ºC, in the presence and absence of vanadate ... 98

Figure 3.43. – Representation of protein activity of the pellet of the control sample at

Figure 3.44. – Representation of the protein activity of the pellet of the control sample

at 4ºC... 99

Figure 3.45. – Representation of absorbance of the supernatant of the control sample

at 4ºC, in the presence and absence of vanadate ... 100

Figure 3.46. – Representation of absorbance of the pellet of the DDM sample at 4ºC,

in the presence and absence of vanadate ... 101

Figure 3.47. – Representation of the protein activity of the supernatant of the DDM

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the presence and absence of vanadate ... 103

Figure 3.50. – Representation of absorbance of the supernatant of the SMA sample, at

Figure 3.51. – Representation of protein activity of the supernatant of the DDM sample

at 20ºC in the presence and absence of vanadate ... 105

Figure 3.52. – Representation of protein activity of the supernatant of the DDM sample

at 20ºC ... 105

Figure 3.53. – Representation of absorbance of the supernatant of the SMA sample at

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– phosphatidylinositol, PE – phosphatidylethanolamine, CI – cardiolipin and PC –

phosphatidylcholine. The table was adapted from [1] ... 19

Table 2.1. – Values of the refractive increments with concentration used for each copolymer, from [30] ... 33

Table 2.2. – Concentration of each compound present in each sample of the experiment I ... 38

Table 2.3. – Concentration of each compound present in each sample of the experiment II ... 39

Table 2.4. – Concentration of each compound present in each sample of the experiment III ... 39

Table 2.5. – Concentration of each compound present in each sample of the experiment IV ... 40

Table 3.1. – DLS measurement for liposomes of DPPC:DMPS (7:3) ... 48

Table 3.4. – DLS measurement for liposomes of DPPC:DMPG (3:1) ... 51

Table 3.5. – DLS measurement for liposomes of POPC:POPE:Cardiolipin (55:35:10) 52 Table 3.6. – Thermodynamic parameters Tm (transition temperature) and ΔH (enthalpy of transition) for the system DPPC:DMPS (7:3) with SMA 3:1 ... 55

Table 3.7. – Thermodynamic parameters Tm and ΔH for the system DPPC:DMPS (7:3) with SMA 2:1 ... 56

Table 3.8. – Thermodynamic parameters Tm and ΔH for the system DPPC:DMPS (6:4) with SMA 3:1 ... 57

Table 3.9. – Thermodynamic parameters Tm and ΔH for the system DPPC:DMPG (3:1) with SMA 3:1 ... 58

Table 3.10. – DLS measurement for liposomes of POPE:POPG (7:3) ... 59

Table 3.11. – Thermodynamic parameters Tm and ΔH for the system POPE:POPG (7:3) with SMA 3:1 ... 61

Table 3.12. – Values of the protein concentration in each sample for experiments with BSA and CM ... 63

Table 3.13. – Properties measured by DLS, of the samples of experiment I, 2h after the addition of SMA ... 65

Table 3.14. – Properties measured by DLS, of the samples of the experiment I, 24h after the addition of SMA ... 66

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the addition of SMA ... 69

Table 3.17. – Properties measured by DLS, of the samples of experiment II, 2h after

the addition of SMA ... 71

Table 3.18. – Properties measured by DLS, of the samples of experiment II, after the

centrifugation ... 72

Table 3.19. – Values of the protein concentration present in each sample for the

experiment II ... 73

Table 3.20. – Properties measured by DLS, of the samples of experiment III, 2h after

the addition of DDM and SMA ... 74

Table 3.21. – Properties measured by DLS, of the samples of experiment III, 24h after

the addition of DDM and SMA ... 75

Table 3.22. – Properties measured by DLS, of the samples of experiment III, after the

experiment III ... 77

Table 3.24. – Properties measured by DLS, of the samples of experiment IV, 2h after

the samples’ preparation ... 79

Table 3.27. – Properties measured by DLS, of the samples of experiment IV, after

Table 3.28. – Properties measured by DLS, of the samples of experiment V, 2h after

samples’ preparation... 90

Table 3.29. – Properties measured by DLS, of the samples of experiment V, 24h after

samples’ preparation... 92

Table 3.30. – Properties measured by DLS, of the samples of experiment V, after

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CI – Cardiolipin

CM - Crude membranes

DMPG – 1,2-dimyristoyl-sn-glycero-3-phospho-(1'-rac-glycerol) (sodium salt) DMPS – 1,2-dimyristoyl-sn-glycero-3-phospho-L-serine (sodium salt)

DPPC – 1,2-dipalmitoyl-sn-glycero-3-phosphocholine LNPs – Lipid Nanoparticles

LUV – Large Unilamellar Vesicle MLV – Multilamellar Large Vesicle MSP – Membrane scaffold protein NBD – Nucleotide binding domain PC – Phosphatidylcholine

PE – Phosphatidylethanolamine PDB – Protein Data Bank P-gp – P-glycoprotein PI – Phosphatidylinositol

POPC – 1-palmitoyl-2-oleoyl-glycero-3-phosphocholine

POPE – 1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine

POPG – 1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1'-rac-glycerol) (sodium salt) Sf9 – Spodoptera Frugiperda

SMA – Styrene-maleic acid

SMALP – Styrene-maleic acid lipid particle TMD – Transmembrane Domains

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1. Introduction

1.1. The importance of cell membranes and membrane proteins

The cell membrane is a fundamental structure in all organisms, separating the inner content from the outer one, providing protection, shape, control of homeostasis and the inlet and outlet of all types of molecules. Membrane proteins have a very important role in cell membranes. Since the lipid composition of each cell membrane varies, each composition will provide different characteristics such as fluidity/hardness, size, formal charge, etc. Therefore, it is important to know the lipid composition and the membrane characteristics together with membrane proteins.

1.2. Cell membranes’ composition

All living species share the fact that lipids are the basic constituents of cell membranes. These lipids can vary in length, saturation, charge, relative amount, etc. For example, plants’ cell membranes tend to have a higher percentage of unsaturated lipids in their membrane as compared to animals. In the case of bacteria, these do not have cholesterol in their membrane composition, but have instead hopanoids which are sterols somewhat similar to cholesterol.

Another common feature of all biomembranes is the fact that they all have proteins associated to the cell membrane and those have a major role in controlling the inlet and outlet of a variety of compounds, and also providing some stability to the cell membrane.

In the present work we wanted to have information on the composition of the species Spodoptera Frugiperda, where P-gp was previously overexpressed in the membranes used in our experiments. Since the ultimate goal of my project was to extract P-gp from these membranes, with the help of the copolymers, forming lipid nanodiscs, we characterized the formation of nanodiscs from model membranes with lipid compositions similar to the insect’s membrane.

According to Kathrin Marheineke et al [1] the plasmatic membrane composition of the insect studied has a great variety of composition, but on average it contains an

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approximately 32% negative charge. The main lipids are phosphatidylethanolamine (PE), phosphatidylcholine (PC), phosphatidylinositol (PI) and cardiolipin (CI), as can be seen in table 1.1.:

Table 1.1. – Phospholipid composition of the membranes of Spodoptera Frugiperda. PI – phosphatidylinositol, PE –

phosphatidylethanolamine, CI – cardiolipin and PC – phosphatidylcholine. The table was adapted from [1]. Fatty acid Total lipids (w/w%) Phospholipid class (w/w%)

PI PE CI PC 14:0 0.2 0.3 0.2 1.3 0.9 16:0 4.5 3.4 4.0 13.4 9.1 16:1 21.5 12.7 19.8 29.5 17.3 18:0 17.9 27.3 15.9 6.2 13.1 18:1ω9 48.0 44.6 52.9 25.7 46.0 18:1ω7 2.6 2.6 2.8 3.7 3.0 18:2ω6 0.7 0.4 0.3 1.8 1.0 20:0-20:3 3.4 5.6 2.5 6.5 2.9 20:4ω6 0.6 1.2 0.6 0.5 0.3 20:5ω3 0.2 0.3 0.2 0.7 0.1 22:5ω3 0.1 0.3 0.1 1.5 0.5 22:6ω3 0.2 0.4 0.3 7.2 3.5 24:1 0.1 1.0 0.4 2.0 2.1

1.3. Introduction to protein extraction

Our body contains proteins, carbohydrates, fatty acids, water (in a very simplified way), and proteins that play a major role. Further, approximately 20% to 30% of the genetic code encodes for membrane proteins [2-4], and membrane proteins represent a target to up to 70% of targeting agents [5].

Protein extraction has been used for the last 50 years for purposes such as enzyme assays, drugs assays, protein sequencing, structure resolution and spectroscopic studies, in order to deepen our knowledge about proteins and their function. When protein structures are resolved, their information is uploaded in a protein data bank (PDB), so that everyone can have access to it. The problem is that

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only less than 0.5% of all structures from PDB are from membrane proteins. Therefore, there is a need to increase the number of membrane protein structures available, to further understand their role and function. However, the problem is that most membrane proteins are not extracted successfully, using the conventional method (detergents), as often the proteins lose their structure and consequently their function [6]. Thus, new alternatives were tried in order to solve this problem.

1.4. Protein extraction methods

Protein extraction started far back in 1980 with the successful extraction of two proteins, bacteriorhodopsin [7] and porin [8]. These extracted proteins were afterwards crystalized, proving that the extraction method was effective, as seen in figures 1.1. and 1.2. Since then, protein extraction through detergents has been widely used. The use of this method was important not only to extract membrane proteins but also to solubilize and manipulate them for posterior biochemical and biophysical characterization [9] as well as in different assays or further studies in other fields such as molecular biology, bioinorganic chemistry, etc.

Figure 1.1. – Representation of bacteriorhodopsin’s crystals.

Figure from [7].

Figure 1.2. – Representation of porin’s crystals.

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Figure 1.3. – Representation of some extraction systems used for proteins, adapted from [6]. Being a – surfactants, b – amphipols, c – bicell, d – membrane scaffold proteins and e –

copolymers.

As non-bilayer system we have commonly used surfactants’ method (figure

1.3. a), but other methods appeared and showed to be effective, such as methods

using amphipols (figure 1.3. b). Within bilayer systems, we have the use of vesicles (not represented in the figure above) and bicelles (figure 1.3. c), nanodiscs maintained by membrane scaffold proteins (figure 1.3. d) and more recently lipid nanodiscs

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formed by copolymers (figure 1.3. e) [6]. The importance of using bilayer systems is very significant, as the lipid environment helps to maintain the structure and function of the extracted protein [10].

1.4.1. Non-bilayer systems

1.4.1.1. Detergents

This method is based on the use of a detergent, or a mixture of detergents, that spontaneously form micelles, as they are able to “solubilise” the membrane forming surfactant micelles, that may contain some remaining lipids [11]. The formation of micelles happens at a certain concentration, that varies depending on the detergent or mixture of detergents, which monomers are able to assemble spontaneously, when reaching the critical micellar concentration, some of which can surround the membrane protein and, allow an efficient protein extraction. Afterwards a purification method is required, as the extracted micelles include not only the protein of interest but other proteins present in the membrane, as well as lipid assemblies. The use of detergents to extract proteins is the most used method, although having considerable disadvantages – among others – the difficulty of finding a proper detergent mix to extract the protein of interest [11, 12], the poor representation of micelles as the cell membrane mimics [13], and last but not least the fact that often surfactants lead to denaturation of the proteins, with consequent function loss [6].

Detergents that are often used for protein extraction can be non-ionic (e.g. dodecyl maltoside (figure 1.4.)), ionic (e.g. sodium dodecyl sulphate (figure 1.5.)) or zwitterionic (e.g. 3-[(3-cholamidopropyl) dimethylammonio]-1-propanesulfonate) (figure

1.6.)). Often these lead to low membrane solubilisation, since the final percentage of

extracted protein is low in many cases, as well as to problems with ion exchange separation, in the case of the ionic detergents (due to the existence of a charge that needs to be stabilize).

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Figure 1.4. – Molecular representation of dodecyl maltoside.

Figure 1.5. – Molecular representation of sodium dodecyl sulphate.

Figure 1.6. – Molecular representation of 3-[(3-cholamidopropyl) dimethylammonio]-1-propanesulfonate.

1.4.1.2. Amphipols

Amphipols are amphipathic polymers that contain a hydrophilic central part and hydrophobic side chains. They are stable, water soluble [14] and are able to coil around proteins, extracting them and maintaining their folded state [15]. This method of protein extraction reveals to be slightly more efficient than using surfactants since it also extracts a small portion of membrane lipids (annular lipids), maintaining some structural native environment, even when being characterized as a non-bilayer method [16] (figure 1.7.). Amphipols vary from each other depending on the nature of the polymers of both central (hydrophilic) and side (hydrophobic) chains, the length, the polar and apolar content, degree of heterogeneity, among other characteristics [17].

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Figure 1.7. – Representation of (A) annular lipids arrangement (yellow) around a protein representation (green) and (B) non-annular lipids (red) arranged nearby a representation of a protein (green). The surrounding lipids (blue) represent

bulk lipids.

1.4.2. Bilayer systems

1.4.2.1. Bicelles

As mentioned before, the use of bilayer systems is very important in order to maintain the structure and function of the extracted protein. Protein stability is not guaranteed when using non-bilayer systems. One possibility is to use synthetic lipids that form vesicles (figure 1.8.) or bicelles (figure 1.9.) to reconstitute the extracted proteins [6] into nanodiscs with lipid composition similar to the original membrane lipids.

The main difference between bicelles and vesicles is in the structure and composition, as vesicles are spherical structures composed by lipids whereas bicelles have a discoidal structure and their composition is based on a mixture of lipids and detergents. The combination of these two types of molecules shows great potential in achieving a better performance in the extraction of proteins [18]. Bicelles have sizes that range from 5 to 50 nm of diameter, depending manly on the detergent/lipid ratio and lipid composition [19].

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Figure 1.8. – Representation of a vesicle, being the blue forms the lipids in their

constitution.

Figure 1.9. - Representation of a bicell, being the blue spheres the lipids and the white ones the detergents’ heads.

1.4.2.2. Nanodiscs bounded by membrane scaffold proteins

Membrane scaffold proteins (MSP) are proteins derived from human apolipoprotein A-1, engineered to have amphipathic helixes [20]. This amphipathic characteristic is useful to protect the hydrophobic part of the lipids from the surrounding polar solvent. These MSPs form nanodiscs with sizes from 6 to 10 nm (the smaller ones) [21, 22] or from 16 to 17 nm (the larger ones) [23]. Using different MSPs or even a different ratio of protein/lipid may lead to the formation of nanodiscs of different sizes ranging from approximately 13 to 21 nm [24].

This method is a very effective and clean one, as the number of MSPs per nanodisc is usually known and constant, and the nanodisc population is quite homogeneous. Nevertheless it is a very expensive methodology being, therefore, less available and not so commonly used.

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1.4.2.3. Nanodiscs stabilized by copolymers

This method is much more recent than the others mentioned so far, and will be described in more detail, as it is the method used in this work.

Until now, copolymers like SMA (Styrene-Maleic Acid) (2:1), SMA (3:1) and DIBMA (DiIsoButylene-Maleic Acid) have been extensively studied for the past few years.

SMA copolymer is composed by styrene (figure 1.10.) and maleic acid (figure

1.11.). The formation of the SMA copolymer is based in the polymerization and

hydrolysis of maleic anhydride and styrene, as shown in figure 1.12. We used SMA 2:1 and 3:1 mixtures. Using the SMA 2:1 sequence as an example, which has two molecules of styrene and one of maleic acid, see figure 1.13.

Figure 1.10. – Molecular representation of

styrene.

Figure 1.11. – Molecular representation of maleic

acid in the protonated form.

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Figure 1.13. – Representation of the sequence of the SMA copolymer.

The mode of action of SMA is based on the formation of nanodiscs, in which SMA folds around a portion of lipids, being then able to extract portions of the vesicles, as represented in figure 1.14. Studies have shown that the styrene units, of SMA, insert themselves in the lipid membrane between the lipid acyl chains [25].

The Styrene-Maleic Acid Lipid Particles (SMALPs) have sizes ranging from 10 to 40 nm, depending on the lipid composition, as previously reported [26-29]. These SMALPs tend to form easily when in a slightly alkaline pH [30] and their contact with high concentrations (more than 5 mM) of divalent cations (e.g. Ca2+ and Mg2+) can lead to SMALPs rupture, since the SMA copolymer is able to chelate with divalent cations [31]. As these copolymers solubilize the membrane lipids’, a partial native organization of the membrane is preserved, surrounded by the copolymer [25, 32]. In recent years, this copolymer was shown to be quite effective in the formation of SMALPs and further, to be able to effectively extract membrane proteins [31, 33-35].

Figure 1.14. – Representation of the addition of SMA in a lipid suspension and the final result. The copolymer is able to

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Another copolymer was more recently described and used, the DIBMA copolymer (figure 1.16.), composed by maleic acid (figure 1.11.) and diisobutylene (figure 1.15.). It has been shown that DIBMA is able to solubilize membrane lipids and extract membrane proteins, and maintaining the native lipid environment less effectively than when using SMALPs [26, 27].

Furthermore, DIBMA does not interfere with optical spectroscopy, as it does not absorb significantly in the wavelength range of interest, making it very suitable to subsequent spectroscopic studies. Even further, while SMA precipitates at low concentrations of divalent cations, DIBMA is more compatible with higher concentrations of divalent ions (approximately up to a concentration of 20 mM) [26, 27]. DIBMA has shown to be more effective in solubilising phospholipids with longer alkyl chains [26].

Figure 1.15. – Molecular representation

of diisobutylene.

Figure 1.16. – Molecular representation of diisobutylene/maleic

acid.

1.5. P-glycoprotein

The aim of this work is to use a copolymer to extract the P-glycoprotein (P-gp) from the membrane of Spodoptera Frugiperda, where P-gp was previously over-expressed. This protein has a molecular weight of 170 kDa and it is overexpressed in cancer cells [37], being responsible for multidrug resistance to chemotherapeutic drugs, since it easily pumps drugs out of the cells.

The mechanism of action of this protein (an efflux pump) is not completely understood. One of the puzzling properties of this protein is how it can recognize and transport so many different substrates. Furthermore it is known that P-gp plays an important role in the excretion of both xenobiotics and pharmacological agents, regulating cellular and tissue levels of these agents. The kinetics of efflux of P-gp

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suggests that this protein removes drugs before they reach the cytoplasm, making this proteins a “hydrophobic vacuum cleaner” [38].

The multidrug resistance that P-gp over-expression generates, in many cases explains why cancer cells survive in the presence of different cytotoxic agents, since this protein does not allow the entrance of those agents into the cells. Besides this, genetic alterations that may occur in resistance to drugs, can lead to different consequences like affect cell cycle dynamics, susceptibility of cells to apoptosis, expression of other efflux proteins, cellular drug metabolism, intracellular compartmentalization of drugs, or repair of drug-induced damage (usually to DNA) [38]. P-gp presents two subunits, each one having a cluster of 6 transmembrane helixes, forming two transmembrane domains (TMD1 and TMD2) each one connected to one nucleotide binding domain (NBD1 and NBD2) [39] (figure 1.17.) and two ATP binding domains.

The transmembrane domains 5, 6, 11 and 12, of P-gp, and the extracellular loops connecting them, were determined to be the principal sites of drug interaction [40-44], although, it is not known if these binding sites are autonomous or interdependent. Besides this, it is know that the two halves of P-gp have an interaction that is necessary for the proper function of the protein, as well as both nucleotide binding domains are essential for this functioning [45-48].

P-glycoprotein is part of the ABC (ATP Binding Cassette) transporters family, and thus uses ATP for its function. It has been reported that the NBDs are fundamental for proper function of the protein. Furthermore, P-gp presents two ATP binding sites, however, these are not able to hydrolyse ATP simultaneously. P-glycoprotein does not have a specific natural substrate, since it is able to transport a great variety group of substances [49].

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Figure 1.17. – Molecular representation of P-gp and its domains, as well as the representation of the lipid membrane in grey lines. Adapted figure from [39].

1.6. Main goal of present work

The main goal of this study was to try to extract P-gp from the insect cell’s membranes, where it was expressed, by using copolymers.

Before starting this work, preliminary methodological work was performed using model membranes and the copolymers of interest. This was needed to gain experience on the main factors controlling nanodisc formation, such as temperature, incubation time and temperature, characterization of the nanodiscs formed and their time and size stability, etc.

Thereafter, the copolymers were used to extract P-gp from the insect membrane of the specie Spodoptera Frugiperda, in order to establish a protocol for this extraction and purification. Hopefully, a successful extraction will enable future drug binding experiments using the protein in solution, stabilized into lipid nanoparticles (LNPs).

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2. Experimental procedure

2.1. Buffers’ preparation

Different buffers were used throughout the work and were always prepared by dissolution in millipore water:

1. Buffer 1 – This-HCl (Tris(hydroxymethyl)aminomethane

hydrochloride) with a concentration of 45 mM, from Roche Diagnostics GmbH, and 50 mM of KCl (potassium chloride), from MERK with 99.5% of purity. After the preparation, the pH was adjusted to 7.50. This was the buffer used in the first step of the experiments (to be explained later);

2. Buffer 2 - 50 mM of Tris base

(Tris(hydroxymethyl)aminomethane), from Sigma-Aldrich with a purity of 99.9%, 500 mM of NaCl (sodium chloride), from Sigma-Aldrich with a purity of 99.5%, and the pH was adjusted to 8.0. That was the buffer used initially to extract the P-glycoprotein with the copolymer SMA 3:1. Later a similar buffer , but at higher ionic strength was used, as the percentage of nanodiscs formed in these conditions was very low;

3. Solubilisation buffer – 50 mM of Tris-HCl (pH = 7.50), 200 mM

of NaCl, from Sigma-Aldrich with a purity of 99.5%, 5 mM of β-mercaptoethanol, 20 mM of imidazole and 15% (w/v) of glycerol;

4. Storage buffer - Tris-HCl (pH=7.50) 45 mM, D-Mannitol 45 mM,

EGTA 1,8 mM (ethylene glycol-bis(β-aminoethyl ether)-N,N,N′,N′-tetraacetic acid), aprotinin 0,45% (v/v) (a polypeptide), DTT 1,8 mM ((2S,3S)-1,4-Bis-sulfanilbutano-2,3-diol), AEBSF 0,9 mM (4-(2-aminoethyl) benzenesulfonyl fluoride hydrochloride) and glycerol 10% (v/v);

5. 2x buffer – Tris-MES (pH = 6.8) 82.4 mM, KCl 100 mM, sodium

azide 10 mM, EGTA (pH ~ 7) 4 mM, ouabain 2,06 mM, DTT 4 mM and MgCl2

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2.2. Copolymers’ preparation

The used copolymers were SMA 2:1 and SMA 3:1. The copolymers were kindly provided by Sandro Keller to Dra. Regina Adão and Prof. Margarida Bastos, already purified and the value for the change in refractive index with concentration provided.

In all experiments performed with model membranes and shown here, the copolymers were dialyzed according to the following protocol:

a) Approximately 1 mL of the copolymer was placed inside a dialysis membrane Spectra/Por® with a cut-off of 3.5 kg.mol-1

b) The membrane was closed on both sides with plastic clamps and put inside a flask with 1000 mL of Tris-HCl buffer, for 16h

c) After 16h the buffer was exchanged for a new and equal one, leaving the dialysis to continue for 8h more

d) At the end of the new 8h period the copolymer is collected, filtered through a 200 nm filter (from Whatman) and its concentration determined by refractometry – at 20ºC

For the determination of the concentration by refractometry, both the buffer (filtered) and the copolymer (dialyzed and filtered) are measured in the refractometer, and the following equation is used to calculate the concentration:

∆𝑛 = 𝑑𝑛

𝑑𝑐× 𝑐 + 𝑛0

Where Δn is the variation of the refractive index, 𝑑𝑛

𝑑𝑐 is the ratio of change in refractive index to the respective change in concentration, c is the concentration we want to determine and 𝑛₀ is the refractive index of the used buffer. The values used for 𝑑𝑛

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Table 2.1. –Values of the refractive increments with concentration used for each copolymer, from [30].

SMA 2:1 SMA 3:1

𝑑𝑛

𝑑𝑐 / L.mol

-1 _0.53 _0.80

2.3. Preparation of model membranes

As referred to above, a methodological approach for the formation of copolymer/lipid nanodiscs using model membranes (Large Unilamellar Vesicles, LUVs) of different lipid composition was performed. The used lipids were all from Avanti Polar Lipids and were DMPG (1,2-dimyristoyl-sn-glycero-3-phospho-(1'-rac-glycerol) (sodium salt)), DMPS (1,2-dimyristoyl-sn-glycero-3-phospho-L-serine (sodium salt)), DPPC (1,2-dipalmitoyl-sn-glycero-3-phosphocholine), POPC (1-palmitoyl-2-oleoyl-glycero-3-phosphocholine), POPE (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphoethanolamine), POPG (1-palmitoyl-2-oleoyl-sn-glycero-3-phospho-(1'-rac-glycerol) (sodium salt)) and cardiolipin (1',3'-bis[1,2-dimyristoyl-sn-glycero-3-phospho]-glycerol (ammonium salt).

a) _b)

c) d)

e) f)

g)

Figure 2.1. – Molecular representation of the used lipids during experiments, a) DMPG, b) DMPS, c) DPPC, d) POPC,

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The model membranes prepared were formed by lipid mixtures, namely: DPPC:DMPS (7:3), DPPC:DMPS (6:4), DPPC:DMPG (3:1), POPE:POPG (7:3) and POPC:POPE:Cardiolipin (55:35:10). In all cases the liposomes were prepared by the lipid film method, as described below:

The necessary amount of each lipid is weighted and solubilized in a mixture of chloroform/methanol (87/13 (v/v)%). The reason for using this particular mixture of chloroform/methanol to solubilize the lipids is because they form an azeotrope, at this proportion, thus keeping the solvent ratio while evaporating. The mixture is allowed to stand for a period that depends on the number of components – 30 minutes for each lipid existing in the mixture. After this, the solvents are evaporated using a rotating evaporator (from Heidolph) at 40ºC, to form the lipid film. Thereafter, the round bottom flask is left under vacuum, overnight, to remove all traces of organic solvents.

After this, the desired volume of buffer 1 (Tris-HCl (pH = 7.50) 45 mM and 50 mM of KCl), previously kept at a temperature at least 5ºC above the gel to liquid crystalline phase transition temperature for the system under preparation, is added to the lipid film and the round bottom flask is left at a bath at the same temperature during 30 minutes, for complete film hydration. Thereafter several cycles of vortex/warming in the water bath are made to assure that all lipid film is removed from the glass and a MLVs suspension is formed.

Finally, the suspension is transferred to cryo-tubes and 3 freeze/thaw cycles (freezing in liquid N2 followed by warming in the water

bath) are made. To obtain a LUVs suspension from the MLVs, an

extrusion process is performed

(extruder from Lipex Biomembranes, Vancouver, BC, Canada (figure

2.2.)). The MLVs suspension is

passed 3 times through a 100 nm filter (Whatman), and after each

passage a cycle of

Figure 2.2. Photography of the extruder used along the experimental work for the extrusion of model membranes.

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vortex/freeze/thaw is made. Finally, the suspension is passed 10 times through 2 superimposed filters of 100 nm with vortex after each passage. The obtained LUVs are kept at a 4ºC overnight prior to their use. All prepared LUVs suspensions were always used within a maximum period of two weeks.

2.4. Lipid quantification

Along the model membrane experiments, the lipid content was quantified by the Bartlett method [50]. The procedure used is described below:

1 – Prepare test tubes with 0, 100, 150, 200, 250 and 300 μL of a 1mM solution

of K2HPO4 and add millipore water to a total volume of 300 μL, in duplicate to generate

the calibration curve.

2 – Prepare triplicates of the samples of the lipid systems with a concentration

within the range of the calibration curve.

3 – Add 700 μL of HClO4 (perchloric acid) at 70% to each tube and vortex.

Cover the tubes with aluminium foil duly punctured.

4 – The tubes are placed in a block heater at 190ºC for 2h, and then cooled to

room temperature.

5 – Add to each tube 2 mL of 1% (m/v) of ammonium molybdate solution and 2

mL of ascorbic acid 4% (m/v) solution.

6 – Vortex and incubate the tubes at 37ºC for 2h. 7 – Read the absorbance of each sample at 700 nm.

2.5. Preparation of lipid nanodiscs

The first part of the work was performed with model membranes (liposomes) and their mixtures with copolymers in order to acquire practical knowledge on these systems and to try to find the best conditions for the next phase of the work, the extraction of P-gp from insect membranes.

For each system, different volumes of copolymer were added to the LUVs to find out the best copolymer/lipid ratio at which concentration the SMALPs were best formed and maintained. The tested P/L ratios for each lipid system were approximately

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0.100, 0.200, 0.300, 0.500, 1.00, and 2.00. However, not in every lipid system all these proportions were tested. The desired volume of copolymer solution is added to the liposomes, and then incubated for 30 minutes at a temperature above the transition temperature of the lipid system, since the formation of nanodiscs is known to be easier when in the fluid phase [28]. The samples are then left at room temperature for 1h and 30 minutes before the first DLS reading, to evaluate the formation or not of nanodiscs. It is possible to visually and qualitatively inspect nanodiscs formation, as the mixture becomes clear instead of a semi-transparent as a common LUV preparation as shown in figure 2.3.

The liposome and liposome/copolymer mixtures were also studied in the DSC in order to evaluate the interaction and understand the behaviour of the samples.

Figure 2.3. – Comparison between liposome suspension (left) and liposome with intermediate copolymer ratio (right).

This picture shows the difference, in aspect, between a lipid suspension with less copolymer (left picture) and a lipid suspension with more copolymer (right picture). The picture with more copolymer is clearer than the one with less

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2.6. Crude membranes of Spodoptera Frugiperda (Sf9)

For this second phase, every procedure is a little different, since the expected results are not the same as the ones obtained, so, for each set of samples a different procedure was used.

All the measurements made by DLS, referred in the following explanations were made at a temperature of 20ºC.

The stock solution of CM had a protein concentration of approximately 6 μg.μL-1

and were obtained from High five insect cells overexpressing mouse P-gp, [51] and were kindly provided by Suresh Ambudkar (Laboratory of Cell Biology, CCR, National Cancer Research, NIH, Bethesda).

2.6.1. Experiment I – testing the influence of time and pH

In this first approach, storage buffer was mixed with buffer 1 (control sample) and CM were mixed with buffer 1 (CM sample). Both these samples had the same volume proportion (150 μL of storage buffer or CM and 1350 μL of buffer 1). Each of the two samples were then mixed with SMA 3:1 (in buffer 1) at two distinct concentrations or buffer 1, leading to a total of 6 samples: control with high concentration of SMA (I), control with low concentration of SMA (II), control with no SMA (III), CM with high concentration of SMA (IV), CM with low concentration of SMA (V) and CM with no SMA (VI). The final composition of each sample is represented in the following table:

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Table 2.2. – Concentration of each compound present in each sample of the experiment I. Compound Sample Tris-HCl/ mM KCl/ mM D-Mannitol/ mM EGTA/ mM % (v/v) aprotinin DTT/ mM AEBSF/ mM % (w/v) glycerol SMA 3:1/mM I 45 47.5 2.25 0.09 0.0225 0.09 0.045 0.5 0.6 II 0.3 III - IV 0.6 V 0.3 VI -

Since after the measurements made, by DLS, were not the expected ones, the conditions of the samples were altered along the time to a final composition of:

 Control with high concentration of SMA – SMA 3:1 was added to a final concentration of 10 mM

 Control with low concentration of SMA – SMA 3:1 was added to a final concentration of 2 mM and pH = 6.63

 CM with high concentration of SMA – SMA 3:1 was added to a final concentration of 10 mM and pH = 8.00

 CM with low concentration of SMA – SMA 3:1 was added to a final concentration of 2 mM and pH = 6.66

2.6.2. Experiment II – testing the influence of a different buffer

In a second try, CM was added to buffer 2 (50 mM of Tris base (pH = 8.00), 500 mM of NaCl and glycerol at 10% (w/v)) was added and shook with gentle vortex. In this experiment, glycerol was added since the aliquots of CM contain glycerol (to better conservation of the cells’ membranes). Then two samples were prepared from the previous mixture and buffer 1 or SMA 3:1:

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Table 2.3. – Concentration of each compound present in each sample of the experiment II. Compound Sample Tris base/mM NaCl/mM Tris-HCl/mM KCl/mM % (w/v) glycerol SMA 3:1/mM Control 37.2 372 - - 7.4 - SMA 5.3 5.9 2.51

For this experiment, in a further step, samples were centrifuged and then analysed by DLS.

2.6.3. Experiment III

– comparing the extraction ability of the

copolymer and of the detergent DDM

In a third attempt, in an aliquot of CM solubilisation buffer was added. For this experiment, the used conditions were based on the extraction method with DDM, so all the next steps were made at 4ºC [52, 53]. 3 different solutions were prepared:

Table 2.4. – Concentration of each compound present in each sample of the experiment III. Compound Sample Tris-HCl/mM NaCl /mM β-mercaptoethanol/mM Imidazole /mM % (w/v) glycerol % (w/v) DDM SMA 3:1/mM Control 32.5 130 3.25 13 9.75 - - DDM 2.0 - SMA - 4.6

For this experiment, samples were analysed by DLS, centrifuged, and again analysed by DLS.

2.6.4. Experiment IV

– testing the influence of different

temperatures

In a fourth experiment, an aliquot of CM was added to solubilisation buffer, as in the previous experiment, but they were incubated at different temperatures. Different

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samples were prepared with SMA 3:1, DDM or none of these (control) and incubated at 4ºC, 20ºC or 40ºC. The final concentration of each component is represented in the following table:

Table 2.5. – Concentration of each compound present in each sample of the experiment IV. Compound Sample Tris-HCl/mM NaCl /mM β-mercaptoethanol/mM Imidazole/ mM % (w/v) glycerol % (w/v) DDM SMA 3:1/mM Control 4ºC, 20ºC and 40ºC 32.8 131 3.28 13.1 9.83 - - DDM 4ºC, 20ºC and 40ºC 2.1 - SMA 4ºC, 20ºC and 40ºC - 4.83

2.6.5. Experiment V

– repeating experiment IV only at 4ºC and

20ºC

This experiment was performed in the same conditions as the experiment IV, but only at 4ºC and 20ºC, since we noticed a significant precipitation at 40ºC. The samples with DDM, SMA or control were then prepared and the final concentrations are represented by the table 2.5.

2.7. Protein quantification method

The protein quantification method used is based on the solubilisation of the membranes to release the proteins, their precipitation and staining.

Samples with a known concentration of bovine serum albumin (BSA) were prepared in parallel to produce a calibration curve.

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2.7.1 Protein quantification of BSA and CM

– testing the

interference of SMA 3:1 in the protein quantification

method

The main goal of this initial protein quantification was to evaluate the possible interference of the copolymer in the quantification method. For that, samples with BSA or with CM were prepared in the presence and absence of SMA 3:1.

Since the goal of this quantification was to assess if the copolymer would interfere in the quantification, and if the interference depended on copolymer concentration, 6 samples of BSA or CM were prepared:

 2 without SMA

 2 contained a low concentration of SMA  2 contained higher concentration of SMA

Afterwards, the protein quantification protocol was followed, as described below:

1 – Using polystyrene tubes standard BSA or protein samples were added to each tube, making a final volume to 300 μL using millipore water.

2 – 30 μL of 10% of SDS (sodium dodecyl sulphate) – a denaturizing agent – is

added to all tubes and then vortexed. 200 μL of 50% TCA (trichloroacetic acid) – a precipitative agent – is also added to each tube and then vortexed.

3 – Wait for 10 minutes at room temperature.

4 – During this incubation, filters (0.45 μm pore size, 47 mm diameter, nitrocellulose MFTM Membrane filters Ref HAWP04700) are prepared and then placed on top of the filter assembly with vacuum on and water is added on top of the filter.

5 – Using an automatic pipette, the samples are spotted on the filter, under vacuum. Then, 6% of TCA is added to the filter and it is then rinsed with water.

6 – The filters are left drying while 4 petri dishes are prepared containing: 1 –

staining solution, 2 – millipore water, 3 – de-staining solution and 4 – millipore water.

7 – The filter(s) is/are left in the petri dish 1 for 3 minutes, then 1 minute in the petri dish 2, then again 3 more minutes in the petri dish number 3 and, finally 1 minute in the petri dish 4 (figure 2.4.). At the end, the filters are left drying for 15 minutes (figure 2.5.).

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Figure 2.4. – Representation of the disposition of the filters and the petri dishes.

Figure 2.5. – Representation of the filters after step 7.

8 – The stained filter spots are cut and transferred to glass tubes with 2 mL of

elution buffer and then vortexed.

9 – After a waiting period of 12-15 minutes at room temperature, the absorption of the solutions at 630 nm is recorded.

Staining solution – 0.1% of napthol blue black in 225 mL methanol + 50 mL of glacial

acetic acid + 225 mL of water.

De-staining solution – 90 mL of methanol + 2 mL of acetic acid + 8 mL of water Elution buffer – 25 mM of NaOH and 0.05 mM of EDTA in 50% ethanol and 50%

water

2.7.2. Protein quantification of the CMs and the protein

extracted with the detergent DDM

The samples that contained the SMA 3:1 copolymer were not quantified, since the presence of the copolymer was found to interfere with the protein quantification. In all cases, samples with different volumes were prepared in duplicate. After this preparation, the protein quantification protocol described above (2.7.1.) was followed.

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2.8. ATPase assay

The ATPase assay uses three compounds, two modulators, verapamil and tariquidar, and one inhibitor, vanadate.

Verapamil – Is a modulator of P-gp that has the ability to increase the activity of

P-gp, it can also be seen as a competitive inhibitor for other substrates [54].

Tariquidar – Is a modulator of the P-glycoprotein that has the ability to inhibit the

ATPase activity when P-gp is in its native membranes, as reported in [54, 55].

Vanadate – Compound that has the ability to inhibit completely the ATPase

activity of the P-glycoprotein [56].

ATPase activity was quantified only for the samples from experiment V, as they represent the optimized conditions we were able to achieve.

In the following procedure, the ATP, SDS, Pi reagent and ascorbic acid were used for:

 ATP for starting the protein activity, since it is ATP dependent;  SDS to stop the ATP dependent reaction, by denaturing the proteins;

 Pi reagent to react with the inorganic phosphate from the hydrolysis of

ATP;

 Ascorbic acid as a reducing agent that forms a coloured complex for the

absorbance measurement.

The following procedure was followed, with all samples prepared in disposable tubes (Lime glass tubes 14x100 mm, VWR Culture tubes ref 212-0017):

1 – Prepare a vanadate solution (9.6 mg of orthovanadate in 0.5 mL of water), and

incubate it at 100ºC (dry bath) for 10 minutes;

2. – Dilute the previous solution 1:100. Read its absorbance to calculate the

concentration (OD = 2.70 corresponds to [V] = 0.75 mM);

3 – Perform a baseline run with water, and thereafter measure the absorbance of

the diluted solution at 268 nm. Do the measurements in triplicate;

4 – Prepare 400 μL of 10 mM solution of vanadate from the original stock solution

point 1);

5 – Place the solution of verapamil and tariquidar (modulators) in DMSO at room

(44)

6 – Prepare control sample by adding water and 2x buffer;

7 – Prepare a protein mix by adding 2x buffer, water and the sample with protein; 8 – Add the protein mix to the corresponding tube;

9 – Add water or vanadate to each tube and DMSO or modulator to each tube.

Leave the tubes in a water bath at 37ºC;

10 – After an incubation period of 3 minutes ATP is added. ATP should be added to

each tube with 30s interval, with mixing after each addition. Incubate for 20 more minutes in water bath (at 37ºC) (from the timepoint of ATP addition);

11 – Add SDS to each tube (with the same time between tubes and the order used

in 10). Vortex each tube after SDS addition.

12 – Add Pi reagent to each tube, water and ascorbic acid, mix well and let it rest

for 10 minutes.

13 – Measure absorbance at 880 nm.

2.9. Description of the used techniques

2.9.1. Dynamic Light Scattering (DLS)

2.9.1.1. Principles and used procedure

Dynamic Light Scattering (DLS) is a technique that measures the size of particles (in solution) and can provide their size distribution. The equipment derives the size of the particles from their hydrodynamic diameter [57] based on the translational diffusion coefficient and the Stokes-Einstein equation, assuming a spherical shape:

D = 𝑘𝐵T 6πηr

where D is the diffusion constant, kB is the Boltzmann’s constant, T is absolute

temperature, η is the viscosity and r is the radius of the spherical particle.

The formation of the nanodiscs and their size characterization were followed by this technique. Disposable cuvettes are used and the measurements are made at