
==== Front
Sci Rep
Sci Rep
Scientific Reports
2045-2322
Nature Publishing Group UK London

39237581
71600
10.1038/s41598-024-71600-2
Article
Raman imaging unveils heme uptake in endothelial cells
Wajda Aleksandra aleksandra.wajda@uj.edu.pl

1
Dybas Jakub 2
Kachamakova-Trojanowska Neli 3
Pacia Marta Z. 2
Wilkosz Natalia 4
Bułat Katarzyna 4
Chwiej Joanna 4
Marzec Katarzyna M. kmarzec@agh.edu.pl

45
1 https://ror.org/03bqmcz70 grid.5522.0 0000 0001 2337 4740 Faculty of Chemistry, Jagiellonian University, Gronostajowa 2, 30-387 Kraków, Poland
2 https://ror.org/03bqmcz70 grid.5522.0 0000 0001 2337 4740 Jagiellonian Centre for Experimental Therapeutics, Jagiellonian University, Bobrzynskiego 14, 30-348 Kraków, Poland
3 https://ror.org/03bqmcz70 grid.5522.0 0000 0001 2337 4740 Malopolska Centre of Biotechnology, Jagiellonian University, Gronostajowa 7a, 30-387 Kraków, Poland
4 grid.9922.0 0000 0000 9174 1488 Faculty of Physics and Applied Computer Science, AGH University of Krakow, al. A. Mickiewicza 30, 30-059 Kraków, Poland
5 https://ror.org/036f4sz05 grid.512763.4 0000 0004 7933 0669 Łukasiewicz Research Network, Krakow Institute of Technology, 73 Zakopianska St., 30-418 Kraków, Poland
5 9 2024
5 9 2024
2024
14 206845 3 2024
29 8 2024
© The Author(s) 2024
2024
https://creativecommons.org/licenses/by-nc-nd/4.0/ Open Access This article is licensed under a Creative Commons Attribution-NonCommercial-NoDerivatives 4.0 International License, which permits any non-commercial use, sharing, distribution and reproduction in any medium or format, as long as you give appropriate credit to the original author(s) and the source, provide a link to the Creative Commons licence, and indicate if you modified the licensed material. You do not have permission under this licence to share adapted material derived from this article or parts of it. The images or other third party material in this article are included in the article’s Creative Commons licence, unless indicated otherwise in a credit line to the material. If material is not included in the article’s Creative Commons licence and your intended use is not permitted by statutory regulation or exceeds the permitted use, you will need to obtain permission directly from the copyright holder. To view a copy of this licence, visit http://creativecommons.org/licenses/by-nc-nd/4.0/.
Heme released from damaged and senescent red blood cells (RBCs) may contribute to oxidant-mediated cell injury. One of the recently investigated physiological processes, essential in preventing the inflammatory impact of labile heme, is its uptake from the bloodstream by endothelial cells (ECs). In this study, we investigated heme uptake by ECs starting from the model studies on the in vitro cellular level, through the endothelium layer on the ex vivo murine aortic tissues. As the cellular model, Human Aortic Endothelial Cells (HAECs) were chosen, and the concentration of labile heme was adjusted so to avoid the excessive toxic effect of the labile heme. We utilized label-free Raman imaging with two different excitation wavelengths to capture the uptake process in situ and characterize the oxidation state of the iron ion in the intercalated heme. The phenomenon of heme uptake was demonstrated in both, the healthy control C57Bl/6J and FVB animals, as well as in mice with developed atherosclerosis (ApoE/LDLR−/− mice). In the presented work, we presented for the first time Raman-based evidence on the heme uptake process by endothelial cells in both, in vitro and ex vivo systems.

Keywords

Endothelial cells
Heme uptake
Raman imaging
In situ label-free imaging
Subject terms

Biochemistry
Biophysics
Cell biology
Molecular biology
Chemistry
Polish National Science CentreUMO-2020/38/E/ST4/00197 Marzec Katarzyna M. issue-copyright-statement© Springer Nature Limited 2024
==== Body
pmcIntroduction

The endothelium constitutes the inner cellular lining of the blood vessel and is the first layer between the bloodstream and the aorta vessel wall. It is composed of approximately 1–6 × 1013 endothelial cells (ECs) anchored to the basal lamina and creating a single layer of cells1. In the past few decades, the role of ECs changed from a passive barrier preventing the access of the blood cells to the vascular matrix into a vital signaling and secretory agent playing a crucial role in cardiovascular homeostasis1–3. Among other functions, ECs secrete prostacyclin and nitric oxide—two potent vasodilators and antiplatelet agents. These endocrine functions make them responsible for controlling blood fluidity, platelet aggregation, and vascular tone1,3. The size of a single EC varies across the vascular tree and can reach up to 70 µm in length and 30 µm in width1. In the blood vessels, they are oriented along the vessel axis, which minimizes the shear forces.

One of the recently recognized ECs roles is the uptake of the labile heme released from deteriorated heme proteins as a consequence of the damage of the circulating erythrocytes4. Degradation of heme proteins can occur when ferrohemoglobin (Fe2+) is oxidized to ferrihemoglobin (methemoglobin, Fe3+)—the natural process happening during the erythrocyte lifespan on the border between blood and aortic tissue4. Such oxidized hemoglobin releases the free ferric heme (Fe3+) to the bloodstream eventually, which can lead to the amplification of the oxidant-mediated cell injury. In normal conditions, heme-scavenging proteins, such as haptoglobin, hemopexin, and albumin tightly bind the free ferric hemoglobin and heme, respectively. However, ECs compete with these heme-scavenging proteins for the free ferric heme, which due to its hydrophobic properties easily crosses the membrane and enters ECs4–6. In contrast, ferrohemoglobin, metmyoglobin, and cytochrome C (CytC) do not alter ECs integrity, as none of them readily releases heme5.

Heme is a potent proinflammatory agent, which can lead to the activation of polymorphonuclear leukocytes7. Moreover, the labile heme can facilitate the expression of ECs adhesion molecule and lead to inflammation. Oxidative damage is mainly associated with nonenzymatic oxidative degradation or enzymatic heme oxygenaze5,8. In the end, the uptake of labile heme alters endothelium, triggers oxidative injury, and may lead to cellular senescence eventually6,9.

The main goal of this research was to observe and in situ image the heme uptake by ECs with the use of Raman imaging. In our experimental setup, we incubated ECs with Hb lysate to track labile heme accumulation. It has to be highlighted that clear-cut differentiation between Hb and Hb-derived heme cannot be provided with the application of Raman spectroscopy. Therefore, we cannot confirm, that observed signals arise solely from labile heme or from Hb. Nevertheless, according to the literature, accumulation of heme by ECs is more preferential compared to Hb4,10, so we decided to use the term “labile heme/heme” in the whole manuscript11,12. The process of heme intercalation is exclusive to ECs in the specific microregions of the aorta, where it is associated with their oxidant-mediated damage. During our search for the heme uptake in cell culture of ECs and endothelium layer within the aorta, we conducted experiments on the healthy control C57Bl/6J and FVB animals, as well as on mice with developed atherosclerosis (ApoE/LDLR−/− mice)13. In situ Raman imaging of the heme distribution was obtained with different excitation wavelengths due to the similar spectral profiles of Hb and CytC. In the context of cellular studies, wherein CytC is more prevalent, utilizing a 488 nm excitation wavelength yielded superior discrimination between the two heme proteins and afforded a deeper understanding of the iron ion oxidation state14–17. Conversely, the 532 nm excitation proved to be more versatile in ex vivo experiments, where cytochromes had minimal impact on heme identification.

Materials and methods

Isolation of red blood cells (RBCs) and preparation of hemoglobin lysate

Whole blood was diluted 1:1 (v/v) with 0.9% saline and subjected to triple centrifugation (acceleration: 500×g; run time: 10 min; RT) followed each time by the removal of supernatant and buffy coat to prepare the isolated RBCs fraction. After the fourth centrifugation, the purity of the isolated RBC fraction was evaluated with a complete blood count, and the number of white blood cells couldn’t exceed 200/µl to process further with RBCs lysis and Hb isolation. Subsequently, RBCs were diluted 1:5 (v/v) with cold deionized water (4 °C) and then frozen at −20 °C. After thawing, the lysed cells were centrifuged at 3000×g for 30 min (4 °C) to remove cell debris and Hb-containing supernatant was collected. The concentration of Hb lysate was assessed by UV–Vis spectroscopy, and calculated based on the absorbance value read at 415 nm18.

Endothelial cell culture and Hb uptake assay

3.5 × 104 human aortic endothelial cells (HAECs) were seeded on a 24-well tissue culture plate in an EGM-MV2 medium (Promokine) and were cultured under standard conditions of 5% CO2 and 37 °C. After 24 h, cells were treated with different concentrations of Hb obtained from lysis described above. The Hb lysate was added to the culture medium in the appropriate concentration. The control cells were cultured in the same way as the tested samples however instead of Hb lysate an equivalent volume of distilled water was added. After 24 h of incubation with Hb, cells were washed 4 times with PBS, and photos of the cells were taken with Motic AE200 microscope. Subsequently, cells were detached with accutase (Thermofisher Scientific) and stained with DAPI to assess cell viability. Cells were analysed for PE-Texas Red autofluorescence and DAPI staining by flow cytometry (BD Fortessa) and subsequently analysed with FlowJo software.

For Raman measurement, 8 × 104 HAECs were seeded on a calcium fluoride glass slide (CaF2, 25 × 2 mm, Crystran LTD) in 200 µl medium. Then, cells were kept for 15 min to attach to the glass surface. Subsequently, 2.5 ml of culture medium was added and cells were incubated for 24 h under normal culture conditions. On the next day, the medium was gently replaced with medium containing the specified amount of Hb (200 µM or 800 µM), or distilled water for the control cells. After 24 h incubation, the cells were washed 4 times with PBS, fixed in 2.5% glutaraldehyde solution for 5 min, and then washed 3 times with PBS.

UV–Vis absorption spectroscopy

UV–Vis electronic absorption spectra of all the samples presented here were obtained on a Perkin Elmer double-beam spectrophotometer Lambda 950 in the range of 300–700 nm using a cuvette of 1 cm path length.

Raman spectroscopy

Raman imaging of single HAECs was performed on the confocal WITec Alpha300R Raman spectrometer (WITec, Germany) equipped with the air-cooled solid state laser operating at 488 nm and 532 nm excitation wavelength, spectral resolution of ~ 3 cm−1 and a low-noise CCD detector (Andor). Raman spectra were collected by the 60× water immersive objective (Nikon Fluor, NA = 1.0) with laser power at the sample position set between 15–18 mW). Raman images were performed with the sampling density of 0.5 μm in x/y direction, and integration time between 0.3–0.7 s. Raman measurements and data analysis were performed using WITec software (WITec Project Plus). The K-means Cluster (KMC) analysis images were obtained after cosmic spike removal and background subtraction in using polynomial shape correction. All Raman spectra presented in this work were taken from the KMC analysis results and are presented after post-processing (cosmic spike removal, smoothing (5–9) and background subtraction).

Ex vivo—tissue preparation and Raman spectroscopy

Male C57Bl/6J mice (wild type), FVB mice and ApoE/LDLR−/− mice (developed atherosclerosis) at the age of 10–16 weeks were anesthetized by intraperitoneal injection (i.p.) of a mixture consisting of ketamine and xylazine (100 mg ketamine/10 mg xylazine/kg body weight). The chest was opened, and the thoracic aorta was isolated. Then the aorta was cut into pieces across (cross-section) or along (en face aorta) the direction of blood flow, as thoroughly described previously19,20. Rings of the aorta were embedded in the OCT medium (Thermo) and frozen at − 80 °C using Leica CM1920 automatic cryostat (Leica, Wetzlar, Germany). The 5 μm thick cross-section slides were put on microscopic glasses coated with poly-L-lysine. Then, slices of the aorta were fixed for 10 min in 4% buffered formalin (Merck). The fragments of en face aorta were glued to the Cell-Tak-coated calcium fluoride surface and preserved by a 10-min soak in 4% buffered formalin.

The Raman images of cross-section or en face aorta were recorded using a WITec confocal CRM alpha 300 Raman microscope equipped with an air-cooled solid-state laser operating at 488 nm and 532 nm as well as a CCD detector which was cooled to -82ºC. The laser was coupled to the microscope via a single mode optical fiber with a diameter of 50 µm. A dry Olympus MPLAN (1006/0.90NA) objective was used. The scattered radiation was focused onto a multi-mode fiber (50 mm diameter) and monochromator. The power of the laser at the sample position was around 15 or 30 mW for 488 or 532 nm excitation wavelengths, respectively. Images with a spectral resolution of 0.33 µm (for 488 nm) or 0.36 µm (for 532 nm) with integration time for a single spectrum of 0.3 s and a resolution of 3 cm−1 were collected. The monochromators of both spectrometers were calibrated using the Raman scattering line produced by a silicon plate (520.7 cm−1). Depth profiling of the tissue was obtained by multiple imaging of the same area in several layers of the sample. Data matrices were analysed using WITec Project software (background subtraction using a polynomial of degree 3 and the automatic removal of cosmic rays). The analysis of the spectra was supported by a Cluster Analysis (CA) (K-means, Manhattan distance, WITec Project Plus).

Ethical approval

C57Bl/6J, FVB and ApoE/LDLR−/− mice were fed a standard chow diet, given water ad libitum, and kept in pathogen-free conditions (22–25°C, 45–65% humidity, 12 h light/12 h dark cycle). At the age of 10–16 weeks were anesthetized by intraperitoneal injection (i.p.) of a mixture consisting of ketamine and xylazine (100 mg ketamine/10 mg xylazine/kg body weight). The chest was opened, and the thoracic aorta was isolated. Then the aorta was cut into pieces across (cross-section) or along (en face aorta) the direction of blood flow, as thoroughly described previously19,20.

All experiments were conducted according to the Guidelines for Animal Care and Treatment of the European Union and approved by the First Local Ethical Committee on Animal Testing at the Jagiellonian University in Krakow. All experimental procedures involving animals were conducted according to the Guidelines for Animal Care and Treatment of the European Communities and the Guide for the Care and Use of Laboratory Animals, published by the US National Institutes of Health (NIH Publication No. 85-23, revised 1996) and in accordance with ARRIVE guidelines.

Results and discussion

Preliminary in vitro study of the heme uptake by the HAECs

After treatment of HAECs with Hb (200 and 800 µM), shape and pattern of the cells was slightly altered (Fig. 1). In contrast, we did not observe a change in cell morphology between 200 and 800 µM. The endothelial monolayer was not disrupted even with the highest Hb concentration used. The staining with DAPI after 24 h of incubation with Hb showed no difference in the cell viability, meaning that in this time-frame, the used Hb concentration was not toxic to the HAECs. There was only a slight increase in PE-Texas red autofluorescence of the HAECs treated with Hb in comparison to the control ones. This increase was not dependent on the concentration of the Hb used.Fig. 1 Evaluation of Hb effect on HAECs after treatment for 24 h with 200 or 800 µM Hb or a vehicle (control). (A) Representative images of HAECs; (B) viability of HAECs assessed by flow cytometry and presented as percentage of DAPI negative cells (live cells); (C) autofluorescence of the HAEC cells in PE-Texas Red channel, shown as median fluorescence intensity (MFI).

In order to determine which excitation wavelength is the most suitable to track heme uptake by HAECs with Raman spectroscopy, we first measured UV–Vis absorption spectra of Hb and oxidized cytochrome C, which is dominant form of CytC in fixed cells21. As presented in Fig. 2, the application of 532 nm excitation provided a similar level of resonance enhancement in the case of both molecules. Moreover, the same heme-related modes connected with the Q0 band are enhanced in case of both molecules. Therefore, differentiation between Hb and oxidized CytC using 532 nm is challenging. The examples of Raman imaging with this excitation were presented in the Supplementary Materials as Figure S1. Even though we applied a high concentration of Hb (800 µM), the tracking of heme uptake still remained ambiguous due to the similarity of its spectral pattern with cytochrome C. In the case of Hb, v4 mode located around 1585 cm−1 is usually more intense, contrary to v21 mode at around 1310 cm−1, which is more intense for cytochrome C compared to Hb. Therefore, their ratio can be a hint in Hb and CytC differentiation (Fig. S1A), but this change is relatively too small to provide clear-cut definition of Hb- and CytC-rich regions.Fig. 2 UV–Vis absorption spectra of hemoglobin and oxidized cytochrome C presented in the 300–700 nm spectral range with marked excitation wavelengths used in Raman imaging.

In turn, application of 488 nm excitation enabled stronger enhancement of Hb bands in Raman spectra compared to CytC, due to the proximity of the Soret band to 488 nm, what is connected with a strong pre-resonance effect16,22,23. It indicated, that heme encapsulated within cells should be characterized by more intense symmetric modes, e.g., ν4, in the Raman spectra in comparison with CytC. Therefore, this excitation was chosen to reveal cells’ heme uptake and differentiate Hb from CytC in in vitro studies. On the other hand, 532 nm excitation was used to capture this process in aorta, where cytochromes did not impede heme identification.

Heme uptake by the HAECs—Raman imaging

In the Fig. 3 there is presented the scheme of the experiments with heme uptake using Raman imaging.Fig. 3 The scheme of the experiment with heme uptake process.

Figure 4 contains examples of Raman images of representative HAECs with heme accumulated during the uptake process. Next to the visual image, each panel comprises Raman integration and K-means clustering (KMC) images obtained with 488 nm excitation line. The Raman integration images present distribution of organic matter, heme, nucleus and lipids. Similarly, KMC images were constructed with division of the consistent dominant classes which were presented with the corresponding Raman spectra averaged from each class.Fig. 4 Representative examples of heme uptaken by HAECs after treatment with 200 µM Hb concentration (A, B and C panels). Each panel comprises visual and Raman images (×60) of the HAECs showing the distribution of organic matter (integration in the 2820–3050 cm−1 range), heme (integration in the 1360–1390 cm−1 range), nucleus (integration in the 780–800 cm−1 range) and lipids (integration in the 2820–2880 cm−1 range), followed by KMC image with color coding as follows: cytoplasm—grey, heme—red, lipids—orange, nucleus—light green and protein rich cytoplasm—dark green. Raman images were recorded with the use of 488 nm excitation. To each cell component there is presented corresponding average Raman spectrum acquired from KMC.

In agreement with the literature, cellular organic matter was distinguished by the integration of broadband in 2820–3050 cm−1 originating from CH2 and CH3 stretching vibrations, typical for biological samples15,24,25. Heme was recognized by the characteristic band located at around 1375–1378 cm−1, which is assigned to v4 mode—pyrrole half-ring stretching vibration15,26,27. As can be noticed in the KMC images, the red class was visible solely in the cellular interior. Close to the cellular membranes were localized lipid droplets, which were characterized by 2850 cm−1 band in the Raman spectra24,28. Their appearance could indicated cellular senescence induced by heme presence, which lead to oxidative cell injury and decreased cell viability in consequence15. Additionally, in the panel C were differentiated regions rich in proteins, associated with other cellular compartments.

More detailed careful analysis of the spectral pattern of the red classes assigned to heme, revealed bands at 1567 and 750 cm−1, which are ascribed to the labile heme—hematin (Fig. S2). Tracking the position of the v4 mode allows to determine the iron ion oxidation state. In case of a ferrous hemes (Fe2+), the v4 is located at around 1360 cm−1, while for a ferric hemes (Fe3+) it blue-shifted to around 1375–1380 cm−116,26,27. In all recorded spectra assigned to the heme class, the v4 mode was located between 1375–1378 cm−1, what taken together indicated free ferric heme (Fe3+). This is also highlighted by the low intensity of the bands at around 675 and 750 cm−1, what is characteristic for the heme without the globin part, in contrast to ferric Hb where these bands are more prominent. Especially, the lack of the band at 675 cm−1 excludes the assignment of the red class to ferric Hb. (Fig. S2). It is worth to mention, that observation of the heme uptake process in HAECs using Raman imaging and 488 nm excitation was possible even in relatively low Hb concentration (200 µM).

Heme uptake by the ECs in the aorta cross-sections and en face

The search for the heme uptake by the ECs in the aorta was started from representative cross-section through the aorta isolated from wild-type C57Bl/6J and FVB mouse strains (Figs. 5, 6, 7). The Raman integration images presented in Figs. 5, 6, 7 and 8 were constructed by integration of certain spectral ranges and represent organic matter (2800–3050 cm−1, stretching vibrations of the CH2 and CH3 groups), elastin (510–550 cm−1, stretching vibrations of the S–S), and heme (ν37 mode originated from ν(CαCm)as in-plane vibrations)15,29.Fig. 5 Representative example of heme uptaken by ECs in the cross-section aorta isolated from wild type mice. Each panel comprises visual and Raman images (×100) of the aorta cross-sections showing the distribution of organic matter (integration in the 2800–3050 cm−1 range), heme (integration in the 1565–1595 cm−1 range), nucleus (integration in the 780–800 cm−1 range), and elastin (integration in the 510–550 cm−1 range) followed by KMC image with color coding as follows: cytoplasm—grey, heme—red, nucleus –green and elastin—blue. Raman images were recorded with the use of 532 nm excitation. To each cell component there is presented corresponding average Raman spectrum acquired from KMC.

Fig. 6 Representative examples of heme uptaken by ECs in the aorta acquired from FVB mice. Each panel comprises visual and Raman images (×100) of the aorta cross-sections showing the distribution of organic matter (integration in the 2800–3050 cm−1 range), elastin (integration in the 510–550 cm−1 range) and heme (integration in the 1565–1595 cm−1 range). Raman images were recorded with the use of 532 nm excitation.

Fig. 7 Representative example of heme uptaken by ECs in the en face aorta isolated from wild type mice. The next panels show images Raman images (×100) of the en face aorta showing the distribution of organic matter (integration in the 2800–3050 cm−1 range), heme (integration in the 1565–1595 cm−1 range), nucleus (integration in the 780–800 cm−1 range), and elastin (integration in the 510–550 cm−1 range) Raman images were recorded with the use of 532 nm excitation.

Fig. 8 Representative examples of heme uptaken by ECs in the aorta acquired from ApoE/LDLR−/− mice. Each panel comprises visual and Raman images (×100) of the aorta cross-sections showing the distribution of organic matter (integration in the 2800–3050 cm−1 range), elastin (integration in the 510–550 cm−1 range) and heme (integration in the 1565–1595 cm−1 range). Raman images were recorded with the use of 532 nm excitation.

Figure 5 shows an example of heme uptake, which is not limited to the endothelial layer itself, but represents the phenomenon of heme entering the subendothelial layers of the aorta. This is evidenced by the location of the Raman signal coming from heme (Fig. 5. Red class) in relation to the location of subsequent elastin layers (Fig. 5. Blue class). The heme uptake was also evidenced in the aorta cross-section from FVB mice (Fig. 6). This stays in agreement with data from the literature, which describes the phenomenon of heme uptake into smooth muscle cells (particularly evident in Figs. 5 and 7). Excess of labile heme induces a concentration-dependent migration and proliferation of vascular smooth muscle cells (VSMCs), which depends on the production of reactive oxygen species (ROS) derived from NADPH oxidase (NADPHox) activity30. Also, heme causes TNF- and ROS-dependent macrophage cell death with characteristics of programmed necrosis31.

Therefore, it demonstrates that heme acts as a double-edged sword. While under physiological conditions heme is crucial for various biological processes, an excess of labile heme acts as a pro-oxidant, enhancing oxidative stress and inflammation32.

To better document that the presence of the heme signal inside the blood vessel was not a contaminant, Raman-based depth-profiling was performed into the blood vessel (Fig. 7). For this purpose, the blood vessel rings were split-open and measured from the depth to the endothelial side. The signal from heme is visible not only on the surface of the blood vessel, but also deep inside, which indicates active incorporation of heme by the endothelium.

In search of the heme uptake by the ECs in the aorta cross-sections

Heme uptake by the endothelium is not a characteristic of solely healthy endothelium, but also occurs in altered blood vessels. We examined representative cross-sections obtained from the murine model of atherosclerosis (ApoE/LDLR−/− mice; Fig. 8A, B). Accumulation of the heme was especially prominent between the elastin fibers. Amount of the heme accumulated within the elastin layers in the aorta cross-sections obtained from ApoE/LDLR−/− mice (Fig. 8A, B) was relatively higher compared to the C57Bl/6J and FVB controls. However, in order to carry out robust quantitative analyses, it is imperative to expand the measurements to encompass a more extensive cohort of atherosclerotic animals. The current study aims to elucidate the measurement methodology and potential, rather than presenting comparative results between control and atherosclerotic mice models.

Conclusions

The application of Raman imaging allowed to track and monitor the heme uptake process in both, in vitro cellular model, as well as in ex vivo tissue studies. The process was not only restricted to control animals (C57Bl/6J or FVB), but was also revealed in the pathologically altered aortic tissue acquired from atherosclerosis mice (ApoE/LDLR−/−), highlighting the potential for future semiquantitative comparisons. We evidenced advantages of different excitation wavelength application. In case of the cellular studies on HAECs the application of 488 nm excitation occurred to deliver more clear-cut distinguishment between CytC and uptaken heme, whereas 532 nm excitation was more versatile tool in ex vivo tissues imaging. As revealed by resonance Raman imaging with 488 nm, the heme uptaken by HAECs occurred in its ferric state (Fe3+). Moreover, recorded spectral profile was the most similar to hematin, which is free ferric form of heme. The results presented herein, may contribute in further studies on proinflammatory heme properties, which might be a possible mediator of lipids oxidation and subsequent ECs injury in case of different civilization diseases.

Supplementary Information

Supplementary Information.

Supplementary Information

The online version contains supplementary material available at 10.1038/s41598-024-71600-2.

Acknowledgements

This research was funded by the Polish National Science Centre, No. UMO-2020/38/E/ST4/00197. Fig. 3 was made using BioRender.com (licensed to M.Z.P.). We would also like to thank Professor Franczyk-Zarow from the University of Agriculture in Krakow, Poland, for generously providing the murine model of atherosclerosis (ApoE/LDLR−/− mice).

Author contributions

Conceptualization and visualization: A.W.; J.D.; N.K.-T.; M.Z.P. and K.M.M.; methodology: A.W.; J.D.; N.K.-T.; M.Z.P.; J.C. and K.M.M., formal analysis: A.W.; J.D.; K.B., N.W.; N.K.-T.; M.Z.P.; investigation: A.W.; J.D.; N.K.-T.; M.Z.P.; K.B.; N.W.; writing—original draft preparation: A.W.; J.D.; N.K.-T.; M.Z.P. and K.M.M.; writing—review and editing: A.W.; J.D.; N.K.-T.; M.Z.P. and K.M.M.; supervision, A.W. and K.M.M.; project administration, K.M.M.; funding acquisition, K.M.M. All authors have read and agreed to the published version of the manuscript.

Data availability

The authors declare that the data supporting the findings of this study are available within the paper and its Supplementary Information files. Should any raw data files be needed in another format they are available from the corresponding author upon reasonable request.

Competing interests

The authors declare no competing interests.

Publisher's note

Springer Nature remains neutral with regard to jurisdictional claims in published maps and institutional affiliations.
==== Refs
References

1. Félétou M The Endothelium, Part I: Multiple functions of the endothelial cells—Focus on endothelium-derived vasoactive mediators Colloq. Ser. Integr. Syst. Physiol. From Mol. Funct. 2011 3 1 306
Félétou, M. The Endothelium, Part I: Multiple functions of the endothelial cells—Focus on endothelium-derived vasoactive mediators. Colloq. Ser. Integr. Syst. Physiol. From Mol. Funct. 3, 1–306 (2011).
2. Szafraniec E Diversity among endothelial cell lines revealed by Raman and Fourier-transform infrared spectroscopic imaging Analyst 2018 143 4323 4334 10.1039/C8AN00239H 30106072
Szafraniec, E. et al. Diversity among endothelial cell lines revealed by Raman and Fourier-transform infrared spectroscopic imaging. Analyst 143, 4323–4334 (2018).30106072 10.1039/C8AN00239H
3. Pearson JD Normal endothelial cell function Lupus 2000 9 183 188 10.1191/096120300678828299 10805485
Pearson, J. D. Normal endothelial cell function. Lupus 9, 183–188 (2000).10805485 10.1191/096120300678828299
4. Balla J Endothelial-cell heme uptake from heme proteins: Induction of sensitization and desensitization to oxidant damage Proc. Natl. Acad. Sci. U.S.A. 1993 90 9285 9289 10.1073/pnas.90.20.9285 8415693
Balla, J. et al. Endothelial-cell heme uptake from heme proteins: Induction of sensitization and desensitization to oxidant damage. Proc. Natl. Acad. Sci. U.S.A. 90, 9285–9289 (1993).8415693 10.1073/pnas.90.20.9285
5. Ferraro JR Nakamoto K Introductory Raman Spectroscopy 1994 Elsevier Science
Ferraro, J. R. & Nakamoto, K. Introductory Raman Spectroscopy (Elsevier Science, 1994).
6. Balla J Heme, heme oxygenase, and ferritin: How the vascular endothelium survives (and dies) in an iron-rich environment Antioxid. Redox Signal. 2007 9 2119 2137 10.1089/ars.2007.1787 17767398
Balla, J. et al. Heme, heme oxygenase, and ferritin: How the vascular endothelium survives (and dies) in an iron-rich environment. Antioxid. Redox Signal. 9, 2119–2137. 10.1089/ars.2007.1787 (2007).17767398 10.1089/ars.2007.1787
7. Graça-Souza AV Arruda MAB de Freitas MS Barja-Fidalgo C Oliveira PL Neutrophil activation by heme: Implications for inflammatory processes Blood 2002 99 4160 4165 10.1182/blood.V99.11.4160 12010821
Graça-Souza, A. V., Arruda, M. A. B., de Freitas, M. S., Barja-Fidalgo, C. & Oliveira, P. L. Neutrophil activation by heme: Implications for inflammatory processes. Blood 99, 4160–4165 (2002).12010821 10.1182/blood.V99.11.4160
8. Balla G Vercellotti G Eaton JW Jacob HS Heme uptake by endothelium synergizes polymorphonuclear granulocyte-mediated damage Trans. Assoc. Am. Physicians 1990 103 174 179 2132529
Balla, G., Vercellotti, G., Eaton, J. W. & Jacob, H. S. Heme uptake by endothelium synergizes polymorphonuclear granulocyte-mediated damage. Trans. Assoc. Am. Physicians 103, 174–179 (1990).2132529
9. Rich AM Armstrong RS Ellis PJ Lay PA Determination of the Fe-ligand bond lengths and Fe-N-O bond angles in horse heart ferric and ferrous nitrosylmyoglobin using multiple-scattering XAFS analyses J. Am. Chem. Soc 1998 120 10827 10836 10.1021/ja980253g
Rich, A. M., Armstrong, R. S., Ellis, P. J. & Lay, P. A. Determination of the Fe-ligand bond lengths and Fe-N-O bond angles in horse heart ferric and ferrous nitrosylmyoglobin using multiple-scattering XAFS analyses. J. Am. Chem. Soc 120, 10827–10836. 10.1021/ja980253g (1998).10.1021/ja980253g
10. Petrillo S Heme accumulation in endothelial cells impairs angiogenesis by triggering paraptosis Cell Death Differ. 2018 25 573 588 10.1038/s41418-017-0001-7 29229999
Petrillo, S. et al. Heme accumulation in endothelial cells impairs angiogenesis by triggering paraptosis. Cell Death Differ. 25, 573–588 (2018).29229999 10.1038/s41418-017-0001-7
11. Soares MP Bozza MT Red alert: Labile heme is an alarmin Curr. Opin. Immunol. 2016 38 94 100 10.1016/j.coi.2015.11.006 26741528
Soares, M. P. & Bozza, M. T. Red alert: Labile heme is an alarmin. Curr. Opin. Immunol. 38, 94–100 (2016).26741528 10.1016/j.coi.2015.11.006
12. Englert FA Labile heme impairs hepatic microcirculation and promotes hepatic injury Arch. Biochem. Biophys. 2019 672 108075 10.1016/j.abb.2019.108075 31412260
Englert, F. A. et al. Labile heme impairs hepatic microcirculation and promotes hepatic injury. Arch. Biochem. Biophys. 672, 108075 (2019).31412260 10.1016/j.abb.2019.108075
13. Marzec KM Vascular diseases investigated ex vivo by using Raman, FT-IR and complementary methods Pharmacol. Rep. 2015 67 744 750 10.1016/j.pharep.2015.05.001 26321276
Marzec, K. M. et al. Vascular diseases investigated ex vivo by using Raman, FT-IR and complementary methods. Pharmacol. Rep. 67, 744–750 (2015).26321276 10.1016/j.pharep.2015.05.001
14. Wood BR Caspers P Puppels GJ Pandiancherri S McNaughton D Resonance Raman spectroscopy of red blood cells using near-infrared laser excitation Anal. Bioanal. Chem. 2007 387 1691 1703 10.1007/s00216-006-0881-8 17151857
Wood, B. R., Caspers, P., Puppels, G. J., Pandiancherri, S. & McNaughton, D. Resonance Raman spectroscopy of red blood cells using near-infrared laser excitation. Anal. Bioanal. Chem. 387, 1691–1703 (2007).17151857 10.1007/s00216-006-0881-8
15. Dybas J Grosicki M Baranska M Marzec KM Raman imaging of heme metabolism: In situ in macrophages and Kupffer cells Analyst 2018 143 3489 3498 10.1039/C8AN00282G 29951676
Dybas, J., Grosicki, M., Baranska, M. & Marzec, K. M. Raman imaging of heme metabolism: In situ in macrophages and Kupffer cells. Analyst 143, 3489–3498 (2018).29951676 10.1039/C8AN00282G
16. Marzec KM Rygula A Wood BR Chlopicki S Baranska M High-Resolution raman imaging reveals spatial location of heme oxidation sites in single red blood cells of dried smears J. Raman Spectrosc. 2014 46 76 83 10.1002/jrs.4600
Marzec, K. M., Rygula, A., Wood, B. R., Chlopicki, S. & Baranska, M. High-Resolution raman imaging reveals spatial location of heme oxidation sites in single red blood cells of dried smears. J. Raman Spectrosc. 46, 76–83 (2014).10.1002/jrs.4600
17. Dybas J Trends in biomedical analysis of red blood cells—Raman spectroscopy against other spectroscopic, microscopic and classical techniques Trends Anal. Chem. 2022 146 116481 10.1016/j.trac.2021.116481
Dybas, J. et al. Trends in biomedical analysis of red blood cells—Raman spectroscopy against other spectroscopic, microscopic and classical techniques. Trends Anal. Chem. 146, 116481 (2022).10.1016/j.trac.2021.116481
18. Wong SS Schenkel OJ Quantification of plasma hemoglobin in the presence of bilirubin with bilirubin oxidase Ann. Clin. Lab. Sci. 1995 25 247 251 7605107
Wong, S. S. & Schenkel, O. J. Quantification of plasma hemoglobin in the presence of bilirubin with bilirubin oxidase. Ann. Clin. Lab. Sci. 25, 247–251 (1995).7605107
19. Pacia MZ Chorazy N Sternak M Wojnar-Lason K Chlopicki S Vascular lipid droplets formed in response to TNF, hypoxia, or OA: Biochemical composition and prostacyclin generation J. Lipid Res. 2023 64 100355 10.1016/j.jlr.2023.100355 36934842
Pacia, M. Z., Chorazy, N., Sternak, M., Wojnar-Lason, K. & Chlopicki, S. Vascular lipid droplets formed in response to TNF, hypoxia, or OA: Biochemical composition and prostacyclin generation. J. Lipid Res. 64, 100355 (2023).36934842 10.1016/j.jlr.2023.100355
20. Pacia MZ Rac1 regulates lipid droplets formation, nanomechanical, and nanostructural changes induced by TNF in vascular endothelium in the isolated murine aorta Cell. Mol. Life Sci. 2022 79 317 10.1007/s00018-022-04362-7 35622139
Pacia, M. Z. et al. Rac1 regulates lipid droplets formation, nanomechanical, and nanostructural changes induced by TNF in vascular endothelium in the isolated murine aorta. Cell. Mol. Life Sci. 79, 317 (2022).35622139 10.1007/s00018-022-04362-7
21. Okada M Label-free Raman observation of cytochrome c dynamics during apoptosis Proc. Natl. Acad. Sci. U.S.A. 2012 109 28 32 10.1073/pnas.1107524108 22184220
Okada, M. et al. Label-free Raman observation of cytochrome c dynamics during apoptosis. Proc. Natl. Acad. Sci. U.S.A. 109, 28–32 (2012).22184220 10.1073/pnas.1107524108
22. Wood BR Hammer L McNaughton D Resonance Raman spectroscopy provides evidence of heme ordering within the functional erythrocyte Vib. Spectrosc. 2005 38 71 78 10.1016/j.vibspec.2005.02.016
Wood, B. R., Hammer, L. & McNaughton, D. Resonance Raman spectroscopy provides evidence of heme ordering within the functional erythrocyte. Vib. Spectrosc. 38, 71–78 (2005).10.1016/j.vibspec.2005.02.016
23. Atkins CG Buckley K Blades MW Turner RFB Raman spectroscopy of blood and blood components Appl. Spectrosc. 2017 71 767 793 10.1177/0003702816686593 28398071
Atkins, C. G., Buckley, K., Blades, M. W. & Turner, R. F. B. Raman spectroscopy of blood and blood components. Appl. Spectrosc. 71, 767–793 (2017).28398071 10.1177/0003702816686593
24. Dybas J Raman spectroscopy as a sensitive probe of soft tissue composition—Imaging of cross-sections of various organs vs. single spectra of tissue homogenates Trends Anal. Chem. 2016 85 117 127 10.1016/j.trac.2016.08.014
Dybas, J. et al. Raman spectroscopy as a sensitive probe of soft tissue composition—Imaging of cross-sections of various organs vs. single spectra of tissue homogenates. Trends Anal. Chem. 85, 117–127 (2016).10.1016/j.trac.2016.08.014
25. Baranska M Optical Spectroscopy and Computational Methods in Biology and Medicine 2014 Springer Netherlands
Baranska, M. Optical Spectroscopy and Computational Methods in Biology and Medicine (Springer Netherlands, 2014).
26. Dybas J Bokamper MJ Marzec KM Mak PJ Probing the structure-function relationship of hemoglobin in living human red blood cells Spectrochim. Acta Part A Mol. Biomol. Spectrosc. 2020 239 118530 10.1016/j.saa.2020.118530
Dybas, J., Bokamper, M. J., Marzec, K. M. & Mak, P. J. Probing the structure-function relationship of hemoglobin in living human red blood cells. Spectrochim. Acta Part A Mol. Biomol. Spectrosc. 239, 118530 (2020).10.1016/j.saa.2020.118530
27. Spiro TG Strekas TC Resonance Raman spectra of heme proteins. Effects of oxidation and spin state J. Am. Chem. Soc. 1974 96 338 345 10.1021/ja00809a004 4361043
Spiro, T. G. & Strekas, T. C. Resonance Raman spectra of heme proteins. Effects of oxidation and spin state. J. Am. Chem. Soc. 96, 338–345 (1974).4361043 10.1021/ja00809a004
28. Czamara K Raman spectroscopy of lipids: A review J. Raman Spectrosc. 2015 46 4 20 10.1002/jrs.4607
Czamara, K. et al. Raman spectroscopy of lipids: A review. J. Raman Spectrosc. 46, 4–20 (2015).10.1002/jrs.4607
29. Pacia MZ Mateuszuk L Chlopicki S Baranska M Kaczor A Biochemical changes of the endothelium in the murine model of NO-deficient hypertension Analyst 2015 140 2178 2184 10.1039/C4AN01870B 25502217
Pacia, M. Z., Mateuszuk, L., Chlopicki, S., Baranska, M. & Kaczor, A. Biochemical changes of the endothelium in the murine model of NO-deficient hypertension. Analyst 140, 2178–2184 (2015).25502217 10.1039/C4AN01870B
30. Moraes JA Heme modulates smooth muscle cell proliferation and migration via NADPH oxidase: A counter-regulatory role for heme oxygenase system Atherosclerosis 2012 224 394 400 10.1016/j.atherosclerosis.2012.07.043 22954673
Moraes, J. A. et al. Heme modulates smooth muscle cell proliferation and migration via NADPH oxidase: A counter-regulatory role for heme oxygenase system. Atherosclerosis 224, 394–400 (2012).22954673 10.1016/j.atherosclerosis.2012.07.043
31. Fortes GB Heme induces programmed necrosis on macrophages through autocrine TNF and ROS production Blood 2012 119 2368 2375 10.1182/blood-2011-08-375303 22262768
Fortes, G. B. et al. Heme induces programmed necrosis on macrophages through autocrine TNF and ROS production. Blood 119, 2368–2375 (2012).22262768 10.1182/blood-2011-08-375303
32. Wagener FADTG Different faces of the heme-heme oxygenase system in inflammation Pharmacol. Rev. 2003 55 551 571 10.1124/pr.55.3.5 12869663
Wagener, F. A. D. T. G. et al. Different faces of the heme-heme oxygenase system in inflammation. Pharmacol. Rev. 55, 551–571 (2003).12869663 10.1124/pr.55.3.5
